UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Biomimetic studies related to lignin degradation Cui, Futong 1990

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
831-UBC_1991_A1 C84.pdf [ 10.44MB ]
Metadata
JSON: 831-1.0059840.json
JSON-LD: 831-1.0059840-ld.json
RDF/XML (Pretty): 831-1.0059840-rdf.xml
RDF/JSON: 831-1.0059840-rdf.json
Turtle: 831-1.0059840-turtle.txt
N-Triples: 831-1.0059840-rdf-ntriples.txt
Original Record: 831-1.0059840-source.json
Full Text
831-1.0059840-fulltext.txt
Citation
831-1.0059840.ris

Full Text

BIOMTMETIC STUDIES RELATED TO LIGNIN DEGRADATION by FUTONG CUI B. Sc., Nankai University, P.R. China, 1984 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY IN THE FACULTY OF GRADUATE STUDIES DEPARTMENT OF CHEMISTRY WE ACCEPT THIS THESIS AS CONFORMING TO THE REQUIRED STANDARD THE UNIVERSITY OF BRITISH COLUMBIA AUGUST, 1990 © Futong Cui, 1990 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of C"neV* nffif; The University of British Columbia Vancouver, Canada DE-6 (2/88) Abstract Lignin is the second most abundant biopolymer on Earth. It is an amorphous, cross-linked, aromatic polymer composed of phenylpropanoid units. There has been an ever growing interest in the biodegradation of this complex polymer for the last 30 years. White-rot fungi have been found to be an important lignin degraders in the natural environment. With the discovery of two groups of hemoprotein enzymes, lignin peroxidases and manganese(II)-dependent peroxidases, from the lignin degrading culture of a white-rot fungus, Phanerochaete chrysosporium, rapid progress has been made in understanding the mechanism of lignin biodegradation. Synthetic metaUoporphyrins, the iron(III) and manganese(m) complexes of meso-tetra(2,6-dichloro-3-sulfonatophenyl)porphyrin (TDCSPPFeCl and TDCSPPMnCl) and mwo-tetra(2,6-dicMoro-3-sulfonatophenyl)-&-c«tacWoroporphyTin (Cl16TSPPFeCl and Cl16TSPPMnCl), were used in this study to rriimic the functions of the "lignin degrading" enzymes. Factors affecting the catalytic activities of these biomimetic catalysts were studied. TDCSPPFeCl could closely mimic lignin peroxidase in the degradation of a number of lignin model compounds, including veratryl alcohol, B-l, B-0;4, B-5, 5-5' biphenyl, phenylpropane, and phenylpropene model compounds. The reactions catalyzed by TDCSPPFeCl include benzyl alcohol oxidation, 0,-0, side chain cleavage, demethoxylation, aromatic ring cleavage, benzylic methylene hydroxylation, and C 0 - C B double bond hydroxylation (glycol formation). Novel solvent incorporated compounds isolated from the oxidation of veratryl alcohol give insights about the site of attack of substrate cation radical by solvent molecules. The isolation of a solvent incorporated product from the oxidation of a phenylpropene model compound suggests a cation radical mechanism for the oxidation of this lignin substructure. The formation of ii a number of direct aromatic ring cleavage products during the oxidation of some model compounds supports the previously proposed mechanism of aromatic ring cleavage. TDCSPPFeCl was also able to catalyze the oxidation of environmental pollutants such as pyrene and 2,4,6-trichlorophenol. Veratryl alcohol and manganese(U)-complexes have been suggested to function as redox mediators for lignin biodegradation. Evidence has been provided to demonstrate their mediating power during electrochemical and biomimetic degradation of lignin model compounds. In addition to the mechanistic information obtained, the successful oxidation of the model compounds suggests that metaUoporphyrins can be important catalysts for the pulp and paper industry and for pollution control. * • • m Table of Contents Abstract i i Table of Contents iv List of Tables x List of Figures xi List of Schemes xx List of Abbreviations xxi List of identification of compounds and structures xxiii Acknowledgements xxviii Chapter 1 Introduction 1 1.1 Structure of lignin 1 1.2 The importance of lignin biodegradation studies 4 1.3 Lignin degrading microorganisms 7 1.3.1 Bacteria 7 1.3.2 Fungi 7 1.4 Lignin degrading enzymes 9 1.5 Reactions catalyzed by lignin peroxidase (LiPl) 14 1.5.1 The mechanism of lignin model compound degradation by L i P l 15 1.5.2 Reactions catalyzed by lignin peroxidase 16 A C 0-CB cleavage 16 iv B B-O-4 bond cleavage and dealkoxylation reactions 19 C Aromatic ring cleavage reactions 19 D Benzylic alcohol oxidations 22 E Glycol formation from C 0 - C B double bonds 25 F. Hydroxylation of benzylic methylene groups 26 G. Degradation (oxidation) of lignin polymers 28 1.6 Biomimetic approaches in lignin biodegradation 29 1.6.1 Degradation of lignin model compounds by iron porphyrins 30 1.6.2 Oxidation of lignin model compounds by inorganic one-electron oxidants 32 1.6.3 Electrochemical oxidation of lignin model compounds 33 1.7 The purpose of this study 34 References for chapter 1 37 Chapter 2 Metalloporphyrins as lignin peroxidase models 45 2.1 Results and discussion 45 2.1.1 Stability of the metalloporphyrins towards excess oxidants 45 2.1.2 Oxidized intermediates of the metalloporphyrins 53 2.1.2.1 Oxidized intermediates of TDCSPPMnCl 55 v A Oxidation of TDCSPPMnCl by sodium hypochlorite 55 B Oxidation of TDCSPPMnCl by H202. t-BuOOH, and mCPBA 58 2.1.2.2 Oxidized intermediates of Cl16TSPPMnCl 61 A Oxidation of Cl16TSPPMnCl by NaCIO 61 B Oxidation of Cl16TSPPMnCl by mCPBA 73 2.1.2.3 Oxidized intermediates of TDCSPPFeCl 75 2.1.2.4 Oxidized intermediates of Cl16TSPPFeCl 75 2.1.3 Factors affecting the catalytic activity of the metalloporphyrins 81 2.1.3.1 pH of the solvent 82 A TDCSPPFeCl as catalyst 82 B Cl16TSPPFeCl as catalyst 92 C TDCSPPMnCl as catalyst 98 D Cl16TSPPMnCl as catalyst 98 2.1.3.2 Oxidant 109 2.1.3.3 The effect of manganese complexes 111 2.1.3.4 The effect of imidazole 116 2.2 Conclusions 120 vi 2.3 Experimental 2.3.1 General 2.3.2 The stability of the metalloporphyrins towards excess oxidants 2.3.3 The oxidized intermediates of the metalloporphyrins 2.3.4 Factors affecting the catalytic activity of the metalloporphyrins References for chapter 2 Chapter 3 Oxidation of lignin model compounds catalyzed by meso-tetra(2,6-dichIoro-3-sulfonatophenyl)porphyrin iron chloride (TDCSPPFeCl) 131 3.1 Results and discussion 131 3.1.1 Oxidation of veratryl alcohol and veratryl acetate 132 3.1.2 Oxidation of phenylpropane model compounds 141 3.1.3 Oxidation of phenylpropene model compounds 145 3.1.4 Oxidation of diarylpropane model compounds 147 3.1.5 Oxidation of arylglycerol-B-aryl ether (B-O-4) model compounds 150 3.1.6 Oxidation of phenylcoumaran (6-5) model compounds 151 3.1.7 Oxidation of 5-5' biphenyl model compounds 154 121 121 124 125 126 128 vii 3.1.8 Oxidation of pyrene 157 3.1.9 Oxidation of 2,4,6-trichlorophenol 158 3.2 Conclusions 159 3.3 Experimental 160 3.3.1 General 160 3.3.2 Syntheses of lignin model compounds 161 3.3.3 The oxidation of the lignin model and related compounds 163 3.3.4 .Characterization of compounds 167 References for chapter 3 189 Chapter 4 Redox mediation in lignin biodegradation 194 4.1 Manganese complexes as mediators in lignin biodegradation 4.1.1 Results 4.1.1.1 Cyclic voltammetry (CV) of manganese complexes 4.1.1.2 Manganese mediated decolorization of Poly B-411 4.1.2 Discussion 4.2 Veratryl alcohol as a possible mediator in lignin biodegradation 4.2.1 Results and discussion 4.3 Experimental 195 196 196 199 208 210 211 217 viii References for chapter 4 221 Chapter 5 Summary 224 Appendices 234 Appendix 1 The oxidation of Lignin Indulin AT 234 Introduction 234 Results and discussion 235 Experimental 240 Appendix 2 Metallophthalocyanines as possible lignin peroxidase models 242 Introduction 242 Results and discussion 243 Conclusions 248 Experimental 270 References for appendices 273 ix List of Tables Table 2.1 Optimum pH for the oxidation of veratryl alcohol by various metalloporphyrin/oxidant 107 4.1 Cyclic voltammetry of manganese sulfate (10 m M solution, pH 4.5) in the presence of various ligands 197 4.2 Comparison of the optimum concentrations of carboxylic acids in the electrochemical decolorization of Poly B-411 and those in the MnP enzyme system 199 4.3 The oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA at various pH values (yield based on substrate) 216 A - l Yield for the oxidation of veratryl alcohol by TSPCMnCl and TSPCFeCl under various conditions 235 A-2 The effect of imidazole on the oxidation of veratryl alcohol by TSPCMnCl and H 2 0 2 at pH 7 234 x List of Figures Figure 1.1 The structure of conifer lignin proposed by Adler 3 1.2 The structure of hardwood (beech) lignin proposed by Nimz 4 1.3 The main substructures of spruce lignin 5 1.4 Proposed catalytic cycle of lignin peroxidase 12 1.5 Proposed catalytic cycle of manganese-dependent peroxidase 13 1.6 The mechanism of C 0 -C f l cleavage of B- l model compounds 17 1.7 The mechanism of C a - C B cleavage of B-O-4 model compounds 18 1.8 The mechanism of B-O-4 cleavage of B-O-4 model compounds 20 1.9 Possible mechanism of demethoxylation of lignin model compounds 21 1.10 Proposed mechanism of aromatic ring cleavage of veratryl alcohol 23 1.11 Possible mechanism of Ca-oxidation of lignin model compounds 24 1.12 Possible mechanism of Ca-hydroxylation of lignin model compounds 27 1.13 The structure of some metalloporphyrins 31 2.1 The relative stability of metalloporphyrins (7.5x10"* M) towards 100 fold excess of hydrogen peroxide (7.5x10"* M) at pH 3 (0.1 M phosphate buffer) in the presence of veratryl alcohol (2.5xl0"3 M) at room temperature 47 2.2 The relative stability of metalloporphyrins (7.5x10"* M) towards 100 fold excess of hydrogen peroxide (7.5x10^ M) at pH 10 (0.1 M phosphate buffer) in the presence of veratryl xi alcohol (2.5xl0"3 M) at room temperature 49 2.3 The stability of TDCSPPFeCl (1x10 s M) towards mCPBA (1x10^ M) in 0.1 M phosphate buffer at room temperature in the absence of veratryl alcohol 50 2.4 The stability of TDCSPPFeCl ( l x l O 5 M) towards hydrogen peroxide (lxlO -* M) in 0.1 M phosphate buffer at room temperature in the absence of veratryl alcohol 51 2.5 The stability of Cl 1 6 TSPPFeCl towards t-BuOOH and H 2 0 2 at pH 3 in the presence of veratryl alcohol at room temperature 52 2.6 UV-vis spectral changes when veratryl alcohol was added to the oxidized intermediate 16 ( T D C S P P M n 1 ^ , X^ 432 nm) 56 2.7 The decay of the oxidized intermediate 17 (TDCSPPMn v=0, 446 nm) to give intermediate 16 (X to„ 432 nm) and the manganese(UI) porphyrin (X^ 472 nm) 59 2.8 The UV-vis spectra of TDCSPPMnCl (X^ 472 nm), T D C S P P M n ^ O (16, 432 nm), and TDCSPPMn v =0 (17, K~ 446 nm) 60 2.9 The conversion of Cl 1 6 TSPPMnCl (X^ 482 nm) to the intermediate 18 (Cl 1 6 TSPPMn n , 452 nm) after veratryl alcohol oxidation stopped 63 2.10 The conversion of the oxidized intermediate 19 (Xma 428 nm) to intermediate 20 (X t a„ 452 nm) upon addition of veratryl alcohol 64 2.11 The reduction of intermediate 20 (X^ 452 nm) to Cl 1 6 TSPPMnCl (X^ 482 nm) by veratryl alcohol 65 2.12 The reduction of intermediate 20 (X^ 452 nm) to Cl 1 6 TSPPMnCl (X^ 482 nm) by veratryl alcohol at pH 10 66 xii 2.13 The UV-vis spectra of intermediate 18 (dash-dotted line) and 20 (continuous line) 68 2.14 The UV-vis spectra of Cl 1 6 TSPPMnCl ( X ^ 482 nm), Cl 1 6 TSPPMn v =0 (19, X t a„ 428 nm), and Cl 1 6 TSPPMn I V =0 (20, X ^ 452 nm) 69 2.15 The oxidation of Cli 6 TSPPMnCl by mCPBA at pH 2 74 2.16 The oxidation of TDCSPPFeCl ( X ^ 424 nm) to the X ^ 432 nm intermediate by NaCIO at pH 10 76 2.17 The oxidation of Cl 1 6 TSPPFeCl ( X ^ 422 nm) to the X ^ 448 nm intermediate by NaCIO at pH 12 78 2.18 The UV-vis spectra of Cl 1 6 TSPPFeCl ( X ^ 422 nm) and Cl 1 6 TSPPFe n -0 2 (k^ 446 nm) 79 2.19 The reaction of CO and Cl 1 6 TSPPFe n -0 2 formed after the oxidation of veratryl alcohol to give Cl 1 6 TSPPFe n -CO ( X ^ 440 nm) 80 2.20 The oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA at various pH values at room temperature 83 2.21 The oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA at pH 5 at room temperature, the same amount of mCPBA was added at time = 0, and time = 1 minute. 84 2.22 The oxidation of veratryl alcohol by TDCSPPFeCl and NaCIO at various pH values at room temperature 85 2.23 The oxidation of veratryl alcohol by TDCSPPFeCl and NaCIO at pH 6 at room temperature, NaCIO added at time = 0 and time = 90 seconds 86 2.24 The oxidation of veratryl alcohol by t-BuOOH catalyzed by TDCSPPFeCl at various pH values at room temperature 87 xiii The oxidation of veratryl alcohol by TDCSPPFeOH and t-BuOOH in pH 3 phosphate buffer containing various concentrations of sodium chloride The oxidation of veratryl alcohol by TDCSPPFeOH and t-BuOOH in pH 3 buffer containing sodium sulfate and sodium chloride The oxidation of veratryl alcohol by TDCSPPFeOH and t-BuOOH in pH 3 phosphate buffer of various concentrations The oxidation of veratryl alcohol by TDCSPPFeCl and hydrogen peroxide at various pH values at room temperature The oxidation of veratryl alcohol by Cl 1 6 TSPPFeCl and mCPBA at various pH values at room temperature The oxidation of veratryl alcohol by Cl 1 6 TSPPFeCl and mCPBA at pH 5 at room temperature, mCPBA added at time = 0 and time = 90 seconds The oxidation of veratryl alcohol by Cl 1 6 TSPPFeCl and t-BuOOH at various pH values at room temperature The oxidation of veratryl alcohol by Clj 6TSPPFeCl and NaCIO at various pH values at room temperature The oxidation of veratryl alcohol by TDCSPPMnCl and mCPBA at various pH values at room temperature The oxidation of veratryl alcohol by TDCSPPMnCl and hydrogen peroxide at pH 10 and pH 9.2 at room temperature The oxidation of veratryl alcohol by TDCSPPMnCl and NaCIO at various pH values at room temperature xiv 2.36 The oxidation of veratryl alcohol by TDCSPPMnCl and mCPBA at pH 5 at room temperature, mCPBA added at time = 0 and time = 240 seconds 102 2.37 The oxidation of veratryl alcohol by Cli 6 TSPPMnCl and NaCIO at various pH values at room temperature 103 2.38 The oxidation of veratryl alcohol by Cl 1 6 TSPPMnCl and hydrogen peroxide at pH 8, 9.2 and 10 at room temperature 104 2.39 The oxidation of veratryl alcohol by Cl 1 6 TSPPMnCl and mCPBA at various pH values at room temperature 105 2.40 The effect of manganese(U)-lactate on the oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA, 1, pH 5 in the presence of M n 2 + ; 2, pH 6 in the presence of M n 2 + ; 3, pH 5 in the absence of M n 2 + ; 4, pH 6 in the absence of M n 2 + 114 2.41 The effect of manganese(JJ)-lactate on the oxidation of veratryl alcohol by Cl 1 6 TSPPMnCl and mCPBA, 1, pH 5 in the presence of Mn 2 + ; 2, pH 5 in the absence of Mn 2 + ; 3, pH 6 in the presence of M n 2 + ; 4, pH 6 in the absence of M n 2 + 115 2.42 The effect of imidazole on the oxidation of veratryl alcohol by hydrogen peroxide and Cl 1 6 TSPPMnCl at pH 9.2, Imidazole/ Cl 1 6 TSPPMnCl (molar ratio): 1, 0; 2, 50; 3, 100; 4, 200 118 2.43 The effect of imidazole on the oxidation of veratryl alcohol by hydrogen peroxide and Cl 1 6 TSPPMnCl at pH 10, Imidazole/ Cl 1 6 TSPPMnCl (molar ratio): 1, 0; 2, 100; 3, 200 119 3.1 The structure of TDCSPPFeCl 131 xv 3.2 The oxidation of veratryl alcohol by lignin peroxidase or biomimetic systems 134 3.3 Proposed mechanism for the oxidation of veratryl alcohol 135 3.4 The oxidation of veratryl acetate in pH 3 aqueous buffer by TDCSPPFeCl and mCPBA 136 3.5 Possible mechanism for the formation of compound 36 137 3.6 The oxidation of veratryl alcohol by TDCSPPFeCl/mCPBA in methanol 138 3.7 Possible mechanism for the formation of compound 37 140 3.8 The oxidation of l-(4-ethoxy-3-methoxyphenyl)propane (44) by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile 143 3.9 Proposed mechanism for alkyl-phenyl cleavage of phenolic model compounds 143 3.10 Possible mechanism for the formation of compound 48 144 3.11 The oxidation of l-(4-ethoxy-3-methoxyphenyl)-l-propene (52) by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile 146 3.12 Possible mechanism for the oxidation of l-(4-ethoxy-3-methoxyphenyl)propene (52) 148 3.13 The oxidation of l-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl) propane- 1,3-diol (55) by TDCSPPFeCl and t-BuOOH 149 3.14 The oxidation of 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether (62) by TDCSPPFeCl and t-BuOOH 151 3.15 The oxidation of 4-O-ethyldehydrodiisoeugenol (64) by TDCSPPFeCl and t-BuOOH 153 3.16 The oxidation of 4,4'-cUethyldehydrodivanillin (74) by xvi TDCSPPFeCl and t-BuOOH 156 3.17 The oxidation of pyrene by TDCSPPFeCl and mCPBA 157 3.18 The oxidation of 2,4,6-trichlorophenol by TDCSPPFeCl and mCPBA 158 4.1 The electrochemical decolorization of Poly B-411 in 0.1 M acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of (L)-lactic acid (mM), 1, 0; 2, 10; 3, 20; 4, 30; 5, 50; 6, 60; 7, 70 200 4.2 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of malic acid (mM), 0, 1, 10; 2, 15; 3, 5 202 4.3 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of (D)-tartaric acid (mM), 1, 10; 2, 20; 3, 25 203 4.4 The electrochemical decolorization of Poly B-411 in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of citric acid (mM), 1, 10; 2, 20; 3, 5 204 4.5 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of malic acid (mM), 1, 10; 2, 50, 3, 60; 4, 40 205 4.6 The electrochemical decolorization of Poly B-411 (0.02%) xvii in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various carboxylic acids at their optimum concentrations, 1, no ligand; 2, 5 m M citric acid; 3, 20 m M (D)-tartaric acid; 4, 60 m M (L)-lactic acid; 5, 10 m M malic acid; 6, 50 m M malonic acid 206 4.7 The decolorization of Poly B-411 (0.005% in 25 mL 0.1 M pH 4.5 acetate buffer) by TDCSPPFeCl ( l x K r 8 mole) and mCPBA (2.5xl0"5 mole) in the presence of manganese sulfate (2.5xl0 5 mole) and lactic acid (1.5x10^ mole), 1, without TDCSPPFeCl 2, without manganese sulfate ; 3, complete reaction 207 4.8 The oxidation of anisyl alcohol (3x10 s mole in 6 mL pH 3 0.1 M phosphate buffer) by TDCSPPFeCl (5x10"* mole) and mCPBA (3xl0' 5 mole) in the presence of veratryl alcohol 211 4.9 The oxidation of anisyl alcohol (3x l0 5 mole in 6 mL pH 3 0.1 M phosphate buffer) by TDCSPPFeCl (5X10"8 mole) and /nCPBA (3xl0' 5 mole) in the presence of 1,4-dimethoxybenzene 212 4.10 The electrochemical decolorization of Poly B-411 (0.005%) in 0.1 M pH 3 phosphate buffer at room temperature in the presence of various amounts of veratryl alcohol (mM), 1, 0; 2, 0.1; 3, 1; 4, 10 214 A - l Difference spectra and reduced-difference spectra of Lignin Indulin A T treated with Cl 1 6 TSPPMnCl and t-BuOOH at pH 2 226 A-2 Difference and reduced-difference spectra of Lignin Indulin A T treated with Cl 1 6 TSPPMnCl and H 2 0 2 at pH 2 227 xviii Difference and reduced-difference spectra of Lignin Indulin A T treated with Cl 1 6 TSPPMnCl and t-BuOOH at pH 10 Difference and reduced-difference spectra of Lignin Indulin A T treated with Cl, 6 TSPPMnCl and H A at pH 10 The Oxidation of 62 by TSPCMnCl (or TSPCFeCl) and mCPBA in aqueous acetonitrile (pH 3) The oxidation of 55 by TSPCMnCl (or TSPCFeCl) and mCPBA in aqueous acetonitrile UV-vis spectral changes of TSPCMnCl and TSPCFeCl QxlO" 7 mole in 3 mL 0.1 M phosphate buffer) upon addition of one equivalent of oxidant ( lxlO' 7 mole) in the presence of veratryl alcohol (5x10"* mole) over a time period of 2 minutes, 10 seconds between scan xix List of schemes Scheme 1.1 The biosynthesis of lignin 1 1.2 C a - C 6 double bond hydroxylation 26 2.1 The redox cycle of TDCSPPMnCl 58 2.2 The redox cycle of Cl 1 6 TSPPMnCl 61 2.3 The effect of superoxide on the catalytic cycle of Cl 1 6 TSPPMnCl during the oxidation of veratryl alcohol 112 3.1 The hydrolysis of compound 37 139 3.2 The biodegradation of dehydrodivanillic acid 155 xx ABTS mCBA Cl 1 6 TSPPFeCl Cl 1 6 TSPPMnCl mCPBA C V DCI DDT D M F ESR FTIR G C GCMS GSH HHIB H P L C HRMS HRP IR LiP MnP M.P. List of Abbreviations 2,2'-azmobis(3-emylbenztMazolme-6^suIfonate) meto-chlorobenzoic acid me.S0-tetra(2,6-dichloro-3-sulfonatophenyl)-B-octacMoropor^^ iron chloride meso-tetra(2,6-dichloro-3-sulfonatophenyl)-B-octachloroporphyrin manganese chloride mete-chloroperbenzoic acid cyclic voltammetry desorption chemical ionization dithiothreitol N,N-dimethylformide electron spin resonance spectroscopy Fourier Transform Infrared spectroscopy gas chromatography gas chromatography-mass spectroscopy glutathione, reduced form hydroxyhexafluoroisobutyrate high performance liquid chromatography high resolution mass spectroscopy horse radish peroxidase infrared spectroscopy lignin peroxidase manganese-dependent peroxidase melting point xxi MS mass spectroscopy MWL milled wood lignin NADPH nicotinamide adenine dinucleotide phosphate, reduced.form NMR nuclear magnetic resonance spectroscopy NOE nuclear overhauser effect PCBs polychlorinated biphenyl's P. chrysosporium Phanerochaete chrysosporium TBAP tetrabutylammonium perchlorate TDCPPFeCl me5o-tetra(2,6-cUcUorophenyl)poiphyrin iron chloride TDCPPFeOH /nejo-tetra(2,6-cUcMOTophenyl)porphyrin iron hydroxide TDCSPPFeCl me5o-tetra(2,6-dlcUoro-3-sulfonatophenyl)porphyrin iron chloride TDCSPPFeOH me5<?-teu*a(2,6^ cUcUoro-3-sulfonatophenyl)porphyrin iron hydroxide TDCSPPMnCl mejt9-tetra(2,6-cUcMorc>3-sulfonatophenyl)porphyrin manganese chloride THF tetrahydrofuran TLC thin layer chromatography TPP meso-tettaphenylporphyrin TPPFeCl /neso-tetraphenylporphyrin iron chloride TSPCCo cobalt tetrasulfonatophthalocyanine TSPCCu copper tetrasulfonatophthalocyanine TSPCFeCl tetrasulfonatophthalocyanine iron chloride TSPCMnCl tetrasulfonatophthalocyanine manganese chloride TSPCNi nickel tetrasulfonatophthalocyanine TSPPFe iron me5o-tetra(4-sulfonatophenyl)poiphyrin UV-vis ultraviolet-visible "Kan absorption maximum in nanometres xxii List of identification of compounds and structures The following numbering system will be used for the compounds and general structures referred to in this thesis. 1 p-coumaryl alcohol (3-(4-hydroxyphenyl)-2-propenol) 2 coniferyl alcohol (3-(4-hydroxy-3-methoxyphenyl)-2-propenol) 3 sinapyl alcohol (3-(4-hyckoxy-3,5-dimethoxyphenyl)-2-propenol) 4 6-0-4 substructure (6-aryl ether) 5 a-O-4 substructure (a-aryl ether) 6 5-5' biphenyl substructure 7 6-1 substructure 8 6-5 (phenylcoumaran) substructure 9 5-0-4 substructure 10 TPPFeCl 11 iron protoporphyrin 12 TDCSPPFeCl 13 TDCSPPMnCl 14 Cl 1 6 TSPPFeCl 15 Cl 1 6 TSPPMnCl 16 TDCSPPMn I V =0 17 TDCSPPMn v =0 18 Cl 1 6 TSPPMn n 19 Cl 1 6 TSPPMn v =0 xxiii 20 C l ^ T S P P M n ^ O 21 oxidized intermediate of TDCSPPFeCl 22 Oxidized intermediate of Cl, 6TSPPFeCl 23 Cl 1 6 TSPPFe n -0 2 24 iron we5o-tetra(2,6-cUmemyl-3-sulfonatophenyl)porphyrin 25 manganese me5o-tetra(2,6-cmiiemyl-3-sulfonatophenyl)porphyrin 26 methyl (Z)-furan-5(2H)-one-3-butenoate 27 methyl (E)-furan-5(2H)-one-3-butenoate 28 methyl (Z)-6-oxo-2H-pyran-3(6H)-ylideneacetate 29 methyl (E)-6-oxo-2H-pyran-3(6H)-ylideneacetate 30 2-hyclroxymethyl-5-methoxy-2,5-cyclohexadiene-1,4-dione 31 2-methoxy-1,4-benzoquinone 32 4,5-dimethoxy- 3,5 -cyclohexadiene-1,2-dione 33 dimethyl 3-hydroxymethyl-2,4-hexadienedioate 34 2-acetoxymethyl-5-methoxy-2,5-cyclohexadiene-1,4-dione 35 3,4-dimethoxyphenol 36 dimethyl 3-acetoxymethyl-2,4-hexadienedioate 37 2-hyclroxymethyl-4,4,5-rrimemoxy-2,5-cyclohexadiene-1 -one 38 2-cUmemoxymemyl-4,4,5-trimethoxy-2,5-cyclohexadiene-1 -one 39 2-acetoxymemyl-4,4,5-rrimethoxy-2,5-cyclohexadiene-1 -one 40 4-ethoxy-2-hych-oxymemyl-4,5-d^ethoxy-2,5-cyclohexadiene-1 -one 41 5-emoxy-2-hych-oxymethyl-4,4-dimethoxy-2,5 -cyclohexadiene-1 -one 42 2-mmethoxymemyl-5-emoxy-3,4-dimethoxy-2,5-cyclohexadiene-1 -one 43 phenylpropane substructure 44 1 -(4-ethoxy-3-methoxyphenyl)propane xxiv 45 (6)-ethyl (l)-methyl 3-propyl-2,4-hexadienedioate 46 2-ethoxy-5-propyl-2,5-cyclohexadiene-1,4-dione 47 l-(4-ethoxy-3-methoxyphenyl)propanol 48 4-hydroxy-2-memoxy-4-propyl-2,5-cyclohexadiene- 1-one 49 2-alkyl-4-hydroxy-3-methoxybenzyl alcohol 50 4-hydroxy-4-(alkylhyfroxymeto^ 1 -one 51 phenylpropene substructure 52 l-(4-ethoxy-3-methoxyphenyl)-l-propene 53 1 -(4-ethoxy-3-methoxyphenyl)propane-1,2-diol 54 4-hydroxy-4- (4-emoxy-3-memoxyphenyl)-3-memylbutyTonitrile 55 1 -(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)propane-1,3-diol 56 4-ethoxy-3-methoxybenzaldehyde 57 4'-methoxy-a-hydroxyacetophenone 58 4'-methoxyphenylglycol 59 4-ethoxy-3-methoxybenzoic acid 60 4-methoxybenzaldehyde 61 4-methoxybenzoic acid 62 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether 63 guaiacol 64 4-O-ethyldehydrodiisoeugenol 65 2,3-dihydro-5-formyl-7-methoxy-3-methyl-2-(4-ethoxy-3-methoxyphenyl)-benzofuran 66 5-carboxy-2,3-dihydro-7-methoxy-3-methyl-2-(4-ethoxy-3-methoxyphenyl)-benzofuran 67 5-carboxy-2,3-dihydro-2-[(6)-ethyl (l)-methyl 2(E)-4-(Z)-2,4-hexadienedioate-3-XXV yl] -7-methoxy-3-methylbenzofuran 68 23-dihydro-5-fonnyl-2-[(6)-ethyl (l)-methyl 2-(E)-4-(Z)-2,4-hexadrenedioate-3-yl]-7-methoxy-3-methylbenzofuran 69 23-dihyclro-5-formyl-2-[(6)-ethyl (l)-methyl 2-(E)-4-(E)-2,4-hexadienedioate-3-yl]-7-methoxy-3-methylbenzofuran 70 2-acetyl-4-formyl-6-methoxyphenyl 4-ethoxy-3-methoxybenzoate 71 guaiacyl substructure 72 dehydrcxiivanillic acid 73 5-carboxyvanillic acid 74 4,4'-cUemyldehydrodivanillin 75 4,4'-cUemoxy-33'-dimemoxy-5-carboxy-5'-formylbiphenyl 76 4,4'-cUemyldehydrodivanillic acid 77 (l)-ethyl (6)-methyl 2-(2-emoxy-3-methoxy-5-formylphenyl)-4-carboxy-2,4-hexadienedioate 78 (l)-ethyl (6)-methyl 2-(2-ethoxy-3-methoxy-5-carboxyphenyl)-4-carboxy-2,4-hexadienedioate 79 pyrene 80 pyrene-1,6-dione 81 pyrene-1,8-dione 82 2,4,6-trichlorophenol 83 2,6-dichloro-1,4-benzoquinone 84 Poly B-411 85 unconjugated guaiacyl substructure 86 conjugated guaiacyl substructure 87 phthalocyanine xxvi 88 general structure of porphyrins 89 TSPCFeCl 90 TSPCMnCl 91 TSPCNi 92 TSPCCu 93 TSPCCo 94 metallo tetrasulfonatophthalocyanine 95 3-(4-ethoxy-3-methoxyphenyl)-2-(2-methoxyphenoxy)-3-oxopropanol 96 3-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)-3-oxopropanol xxvii Acknowledgements I wish to express my sincere thanks to Professor David Dolphin. His guidance and encouragement have been an invaluable support throughout my research. I am indebted to Dr. Tilak Wijesekera for kindly providing samples of meso-tetra(2,6-dichlorophenyl)porphyrin and me,s<9-tetra(2,6-dichlorophenyl)-B-octachloroporphyrin, for his guidance in the syntheses of the metalloporphyrins used in this study, and for reading this thesis. I would also like to thank all the members of Professor Dolphin's research group, past and present, for their invaluable help and discussions. I would like to take this opportunity to express thanks to my wife and my family for their support and understanding. xxviii In memory of my father J Chapter 1 Introduction 1.1 Structure of lignin Lignin is a three-dimensional, amorphous aromatic biopolymer occurring widely in vascular plants. It is among the most complex biopolymers in nature and the understanding of its chemical structure is based on studies of its biosynthesis and degradation. It is now known that lignin is formed in plants at the site of lignification by enzyme mediated dehydrogenative polymerization of three precursors, p-coumaryl alcohol (1), coniferyl alcohol (2) and sinapyl alcohol (3) as shown in Scheme 1.1. The three phenylpropanoid units in lignin are connected randomly by various C-C and C-O Scheme 1.1 The biosynthesis of lignin 1 bonds. The proportions of each phenylpropanoid unit vary with the origin of lignin. Higuchi et aP defined three major groups of lignin by their composition: guaiacyl lignin, guaiacyl-syringyl lignin, and guaiacyl-syringyl-coumaryl lignin. Guaiacyl lignin is composed of coniferyl alcohol units with small amounts of coumaryl and sinapyl alcohol derived units. Guaiacyl-syringyl lignin contains approximately equal amounts of coniferyl alcohol and sinapyl alcohol units and only small amounts of coumaryl alcohol units. Guaiacyl-syringyl-coumaryl lignin is composed of approximately equal amounts of the three cinnamyl alcohol units. Lignin is sometimes also named by the type of plant from which it was prepared. Examples are conifer lignin, softwood lignin, hardwood lignin, and grass lignin etc. It should be noted that the structure of lignin varies with the specific species, age, and probably also the environment of the plant from which the lignin was obtained. As lignin is closely associated with cellulose in plants in the form of lignocellulose, physical (i.e. solvent extraction ), chemical ( i.e. acid treatment), or biological ( i.e. enzyme treatment ) methods have to be used to separate lignin from cellulose. The chemical and physical properties of lignin are inevitably altered to different extents by different isolation procedures. Some methods can isolate only a small portion of lignin which may not be representative of the integral sample. In addition, lignin preparations are likely to be contaroinated by residual cellulose and other components such as proteins, and inorganic compounds. It is almost impossible, therefore, to obtain "natural" lignin sample. The composition, molecular weight, and chemical structure of a specific sample is also affected by its preparative procedure. Strictly speaking, it is impossible to know the exact molecular structure of a lignin molecule. It is possible, however, to roughly represent the structure of lignin by constructing a portion of a lignin molecule and Figure 1.1 shows a scheme proposed by 2 I 9 H 2 ° H t H C = 0 [ C H 2 O H ] I C H 2 O H I ^ H C - O -| f S Ar-"" ' C H 3 ° C H O - S ^ C H 2 0 H JL'€T H C 0 H ? 0 C H J J , P fijii H O H J C - C - C H C O H H O C H , ^ Q C H 3 2 " ^ 0 0 x ' I I H C C H I I C O H C H j O H 0 » H C — - — 6 J " " V S & \ H 0 H ^ C C H j O 1 ^ - ^ O C H 3 H C 0 0 I " J®L H O < £ > C H H ^ O / C H 2 I C H 3 O T O 1 " 1 - 1 - " " m u ^ " V . O C H 3 O C H ^ H J O H j , 0 -O C H H C — 0 H C - O H C H , 0 C H , O H H ^ O H C H 3 0 -— C H H C H C O H A . c=o JL &\ K L v — VSjCHj O H O H [ O - C ] Figure 1.1 The structure of conifer lignin proposed by Adler 1 Adler2 representing conifer lignin. The phenylpropanoid units were connected by linkages which have been shown to occur in lignin. The proportions of various linkages shown approximate those of the lignin sample, though the importance of a few structural types is overemphasized due to the number of units that may be assembled into a scheme of this size. A scheme representing the structure of hardwood (beech) lignin proposed by Nimz 3 is shown in Figure 1.2. From Figures 1.1 and 1.2 it can be seen that the structure of lignin is very complex and this kind of schematic presentation refers only to an "average" lignin. It is easiest to view lignin as composed of various substructures which occur in various proportions in different lignins. Figure 1.3 shows some of the most important substructures in spruce lignin. These include arylglycerol-fJ-aryl ether (B-0-4), noncyclic benzyl aryl ether (a-0-4), biphenyl (5-5'), 1,2-diarylpropane (B-l), and 3 H 2 C O H Figure 1.2 The structure of hardwood (beech) lignin proposed by N i m z 3 phenylcoumaran (B-5) substructures. 1.2 The importance of lignin biodegradation studies Lignin is the second most abundant biopolymer on earth3 and it has been estimated that 25% of the organic compounds produced by photosynthesis is eventually transformed into lignin. It is, however, one of the most persistent naturally occurring compounds and its biodegradation is probably the rate hmiting step in the biospheric carbon-oxygen cycle. In addition, the most abundant biopolymer, cellulose, is protected by lignin in woody plant tissues and the efficient biodegradation of cellulose requires 4 R 7, P-i 8, 0-5 9, 5 - 0 - 4 Figure 1.3 The main substructures of spruce lignin. R = H , O C H 3 or lignin pre-disruption of the protective lignin sheath. The biodegradation of lignin, therefore, is critical for the decomposition of biomass in nature. On the other hand, the undesirable decay of lignin in woody construction materials causes enormous economical damage, and a better understanding of lignin degradation could also provide knowledge about wood preservation. Being a renewable organic material, lignin is also an important natural resource. Billions of tons of lignocellulosic materials are produced annually in the world. Biotechnology holds a great potential to transform lignin into chemical raw materials4"5,6 and animal feed stock.7 Due to its unique molecular structure, however, lignin is difficult to handle by most enzymes and microorganisms. Attempts to develop industrial processes for the conversion of lignin into useful products have been unsuccessful. Biological degradation of lignin into low molecular weight aromatics is probably the bottleneck for the industrial utilization of this abundant valuable resource. In the pulp and paper industry, chemical or mechanical methods are used to remove lignin from the desired cellulose (the pulping process). Chemical pulping gives high quality pulp, but pulp yield is low. The mechanical pulping process gives pulp in high yield without much pollution but the pulp quality is not acceptable in most applications and needs high energy input for its prodution. Pulp bleaching nowadays also depends on chemicals and produces environmentally hazardous chlorinated materials in the process. Biomechanical pulping, using microorganisms or enzymes in combination with mechanical methods as the means of delignification, has been proposed as an alternative pulping process. Thanks to the rapid progress of lignin biodegradation studies, one of the lignin degrading white-rot fungi, Phanerochaete chrysosporium, has been used successfully in the laboratory scale to pre-treat wood chips8 in order to reduce the energy input for the subsequent mechanical pulping and to 6 increase the pulp quality. Microorganism cultures9 and biornimetic catalysts10 have also been used for pulp bleaching on a bench scale. Successful application of biotechnology in this area could revolutionize the pulp and paper industry in the near future. Added to the importance of lignin biodegradation studies is the possible application of lignin degrading systems in the degradation of environmentally persistent pollutants. Chlorinated lignin, polychlorinated biphenyls (PCBs), and polycyclic aromatics are slowly metabolized by ligninolytic P. chrysosporium culture to give carbon dioxide. Dark pulp mill effluent is also decolorized by P. chrysosporium. 1.3 Lignin degrading microorganisms 1.3.1 Bacteria A variety of bacteria, including strains of Norcardia and Streptomyces, have been reported to be able to degrade lignin. 1 1 1 3 Studies using [14C]-lignin specifically labelled at side-chain, aromatic ring, and methoxyls, respectively, showed that all the three structural components are converted to 1 4 C 0 2 to various extents.14 Bacteria, however, seem to play a secondary role in lignin degradation. None of the bacterial strains can cause rapid and extensive degradation of lignin compared to the white-rot fungi. 1.3.2 Fungi Lignin degrading fungi are classified as white-rot (Basidiomycetes), brown-rot (Basidiomycetes), and soft-rot (Ascomycetes) depending on the pattern of their action 7 on wood. Brown-rot fungi mainly decompose the carbohydrate in wood, leaving a modified, brown lignin residue. Analysis of the resulting lignin residue showed that the major consequence of brown-rot attack is demethoxylation of aromatic methoxyl groups and direct hydroxylation of the aromatic nucleus. The carbonyl and carboxylic acid groups also increased after the attack, indicating the oxidative nature of the brown-rot attack.13,16 Studies using [14C]-lignin showed that brown-rot fungi have limited ability to degrade the lignin polymer backbone.17 Soft-rot fungi cause a softening of the surface layer of wood, resulting in significant weight loss at the same time. The soft-rot fungi are able to mineralize all the three structural moieties of lignin (aromatic nucleus, side chain, and aromatic methoxyls) in wood, the polysaccharide fraction, however, seem to be preferentially degraded. Lignin decomposition by soft-rot fungi seems to be slow compared to that by white-rot fungi. The soft-rot fungi, however, have not received extensive study and their potential for lignin degradation is still not clear. White-rot fungi can degrade all the major components of wood and are generally believed to be the major microorganisms responsible for lignin degradation in nature. The polysaccharide and lignin fractions of wood are often degraded simultaneously by white-rot fungi. The relative rates at which cellulose, hemicellulose and lignin are degraded depend on the species of fungus and the culture condition. Species are known which selectively degrade lignin and hemicellulose. Of the many species of lignin degrading white-rot fungus, Phanerochaete chrysosporium is the most widely studied and the best lignin degrader found so far. Culture conditions for lignin biodegradation by P. chrysosporium have been optimized by Kirk and coworkers.18 The lignin degrading enzymes of P. chrysosporium are produced only during secondary metabolism. Veratryl alcohol (3,4-dimethoxybenzyl alcohol), a secondary metabolite of 8 P. chrysosporium, was found to be able to induce the biosynthesis of lignin degrading enzymes.19,20 Studies have shown that added lignin also increases the titers of lignin degrading enzymes.21,22 1.4 Lignin degrading enzymes The complex chemical structure of lignin required that its mechanism of degradation differ from that of other biopolymers, such as cellulose and proteins which are linear molecules composed of repeating units connected by a single, hydrolysable, chemical bond. The heterogeneous nature of lignin determined that its degradation can not be handled by enzymes which have substrate specificity. The subunits of lignin are connected by stable C-C and C-O bonds and are not easily broken down by hydrolysis. The lack of knowledge about lignin degrading enzymes led to the proposal that activated oxygen species, superoxide (02"* ), and hydroxy radical ( HO 1 ) were involved in lignin biodegradation.23 Later studies showed, however, that these species play at most a minor role. 2 4 , 2 5 The first lignin degrading enzyme, lignin peroxidase (LiP), was isolated from P. chrysosporium in 1983.26 ,27 It is a hemoprotein containing one iron protoporphyrin IX (protohemin) per molecule. A manganese-dependent peroxidase (MnP), which is also a hemoprotein, was isolated later from the ligninolytic culture of the same white-rot fungus.28,29 Detailed studies30 of the extracellular enzymes of the P. Chrysosporium ligninolytic culture by analytical electric focusing revealed the presence of twenty-one enzymes, fifteen of them are isoenzymes of the previously isolated lignin peroxidase26,27 and six of them belong to the family of the manganese-dependent peroxidase.28,29 9 The lignin peroxidase isoenzymes31 have similar molecular weights, between 38,000 and 43,000, but different isoelectric points, varying between pi 4.7 and 3.3. Al l the lignin peroxidase are able to oxidize veratryl alcohol to veratraldehyde in the presence of hydrogen peroxide. The isoenzymes, however, have different kinetic properties towards various substrates, suggesting different substrate affinities or active site accessibilities. The similarities and differences of the MnP isoenzymes have not been studied in detail. Preliminary studies30,31 showed that they could not oxidize veratryl alcohol, but could oxidize vanillylacetone and phenol red in the presence of Mn(II) and hydrogen peroxide. The importance of the multiplicity of the lignin degrading enzymes is still not clear. The fact that the lignin peroxidases differ in their kinetic properties31, stabilities32 and in their relative concentrations at different culture times31 indicates that some of them might be more important than others for lignin biodegradation. Genetic studies of the isoenzymes suggested that they may have different functions.33 The abilities of different lignin peroxidase isoenzymes to cleave the various linkages in lignin model compounds have not been demonstrated. Comparison of the reactions catalyzed by the major molecular form of lignin peroxidase, L iP l , and the major isoenzyme of the MnP, named MnPl , using the nomenclature system suggested by Farrell et aP1, seems to suggest that these two groups of enzymes catalyze different reactions. MnPl normally catalyzes the oxidation of phenolic model compounds only,34 though the oxidation of non-phenolic model compounds was also reported recendy.3S While L iP l was reported to be able to oxidize a variety of non-phenolic model compounds and to partially depolymerize lignin, the degradation of phenolic model compounds has not been reported. The fact that lignin peroxidase and manganese-dependent peroxidase appear 10 to reach their maximal activities at different culture times30 also suggests that they have different functions in lignin biodegradation. The major lignin peroxidase isoenzyme, LiPl, has a molecular weight of 42,000 daltons and contains about 13% carbohydrate.36 A single crystal was obtained37 but the crystal structure is still unavailable. The spectral properties of the hydrogen peroxide-oxidized species of LiPl were very similar to those of typical peroxidases.3638 Like horseradish peroxidase (HRP) compounds I and n, the compounds I and II of LiPl contains 2 and 1 oxidizing equivalents, respectively, over the ferric resting state of the enzyme.39 Resonance Raman spectra40*41 showed that the local heme environment of LiPl gready resembles those of other peroxidase. At about 25°C, the ferric iron of LiPl is mainly high spin, pentacoordinate, but the high spin hexacoordinate state dominates at below 2°C. Like horseradish peroxidase, LiPl has a vacant coordination site, trans to the proximal fifth ligand, which is histidine,40'42 at ambient temperature. The Resonance Raman spectra of LiPl compound II are also very similar to that of HRP compound II, indicating a low spin, hexacoordinate Fe^^O structure of LiPl Compound n. LiPl was first thought to be an oxygenase.36 It was found later, however, that oxygen was not consumed in substrate amounts in the oxidation of veratryl alcohol by LiPl and hydrogen peroxide.43 Diarylpropane model compounds can be oxidized by LiPl under both aerobic and anaerobic conditions.44'45 l s O from I V O was found to be incorporated into the products under anaerobic conditions.45 Single-electron transfer agents are able to oxidize lignin model compounds in a manner similarly to LiPl . 4 3 LiPl, therefore, does not catalyze the oxidation of lignin model compounds by oxygen activation. It is a peroxidase rather than an oxygenase and oxidizes lignin model compounds by single-electron oxidations. The catalytic cycle of lignin peroxidase has been proposed and is shown in Figure 1.4.39,46 In the absence of a readily oxidizable 11 substrate, excess hydrogen peroxide converts L iP l to its inactive form, compound in.39'46*47 Superoxide can convert the native enzyme direcdy to compound DJ. (see Figure 1.4) 0 Compound I Lignin H 2 0 2 Resting Enzyme Ligninox Lignin Compound II H 2 0 2 02* Compound III Figure 1.4 Proposed catalytic cycle of lignin peroxidase 39,46 The isoenzyme of MnP, MnPl, has a molecular weight of 46,000 daltons and has a protohemin as its prosthetic group.28 Enzyme activity is stimulated by a-hydroxycarboxylic acids and absolutely depends on Mn(U) for its activity. Maximal enzyme activity appears at pH 4.5. The enzyme shows properties of both an oxidase and peroxidase.48 As an oxidase, it oxidizes NADPH, GSH, and DDT ( dithiothreitol ) to give hydrogen peroxide, suggesting that it might work as a hydrogen peroxide 12 generating enzyme for the lignin degrading enzymes of P. chrysosporium. As a peroxidase, the hydrogen peroxide oxidized MnPl can oxidize Mn(U) to give Mn(IU), which in turn oxidizes organic substrates. In the presence of hydrogen peroxide and Mn(H), MnPl can oxidize a number of phenolic compounds and polymeric dyes.29,48 Electronic absorption, ESR, and resonance Raman spectra studies29,49,50 of MnPl showed that the heme iron in the native enzyme is in the high spin pentacoordinate ferric state with histidine as the fifth ligand. The catalytic cycle of MnP has been proposed50 and is shown is Figure 1.5. The oxidation of the native enzyme by one molecule of Compound III Figure 1.5 Proposed catalytic cycle of manganese-dependent peroxidase1 13 hydrogen peroxide gives MnP compound I, reduction of which by Mn(II) or other one electron acceptors (such as phenols) gives the MnP compound U. The Mn(m) generated can in turn oxidizes phenols and is itself reduced back to give Mn(JJ) again. The Mn(n)/Mn(IU) redox couple thus acts as a mediator for the MnP catalyzed oxidations. The compound n of MnP can be reduced by one equivalent of Mn(H) to the native enzyme. Mn(U) is actually the only substrate that reduces MnP compound U efficiently and thus is necessary to maintain the catalytic cycle of MnP. In the absence of Mn(II), MnP compound II is converted to the compound HI by hydrogen peroxide. Laccase is produced by most white-rot fungi (but not by P. chrysosporium) and has long been the subject of research. The main effect of laccase was found to be polymerization of lignin51,52 and related phenolics.53 Morohoshi et at* isolated three extracellular laccases, laccase I, n, and HI from Corriolus versicolour culture by gel filtration and ion-exchange chromatography. The main function of laccase I is to polymerize lignin while laccase HI can depolymerize lignin.55,56 The electrophoretically homogeneous laccase preparation and laccase IU of Coriolus versicolour were reported to be able to catalyze both C^C^ and phenyl-alkyl cleavage reactions of phenolic B-l and B-O-4 dimeric model compounds.57,58 1.5 Reactions catalyzed by lignin peroxidase (LiPl) Due to the complex structure of lignin and the difficulties in obtaining well defined lignin samples, lignin model compounds, which represent a portion of a structure known to occur in lignin, are often used in lignin biodegradation studies. Although it is still uncertain that an enzyme which can degrade lignin model compounds will degrade lignin in the same way, lignin model compounds are still being used as 14 convenient substrates and have played important roles for the elucidation of the molecular mechanisms of lignin biodegradation. The first lignin degrading enzyme26,27 was actually discovered by using a diarylpropane type model compound. Lignin model compounds containing one, two, three, or even four phenylpropanoid (C6-C3) units have been used and are named as monomeric, dimeric, trimeric, and tetrameric model compounds, respectively. Dimeric model compounds are the most commonly used for mechanistic studies and can be classified into several groups depending on the type of linkages connecting the two phenylpropanoid units. The most frequently used dimeric model compounds are the B-O-4 dimers representing the B-O-4 substructure (4, Figure 1.3), B-l dimers representing substructure 7, B-5 dimers representing the phenylcoumaran substructure 8, and biphenyl model compounds representing the biphenyl substructure 6 (see Figure 1.3). 1.5.1 The mechanism of lignin model compound degradation by LiPl Lignin peroxidase is a heme peroxidase which functions by one electron oxidations. Kersten et al59 first observed, by ESR, the aryl cation radical from the oxidation of alkoxybenzenes by LiP and hydrogen peroxide and suggested that lignin peroxidase degrades lignin through an aryl cation radical intermediate. Schoemaker et af° and Harvey et at* also proposed that lignin peroxidase, once oxidized by hydrogen peroxide, can abstract an electron from the electron rich oxygenated aromatic units of lignin to give cation radicals. Further non-enzymatic fragmentation of the cation radical leads to the degradation of lignin. The single electron oxidation mechanism of lignin model compound degradation by ligninase was demonstrated by Hammel et a/.44 In the oxidation of a B-l model 15 compound by LiP , both C a - and Ce-centered radicals were detected. Evidence was also obtained that the free radicals were intermediates during C a - C B cleavage. Radical coupling products of CB-centered radicals were found under anaerobic conditions and a hydroperoxide derived from the CB-centered radical was found in the presence of oxygen. A cation radical of a C a-carbonyl B-O-4 dimer was observed by ESR. 6 1 Thus, lignin peroxidase degrades lignin model compounds through a cation radical pathway. A l l the known reactions catalyzed by lignin peroxidase can be rationalized by this single electron oxidation mechanism. 1.5.2 Reactions catalyzed by lignin peroxidase A . C a - C B cleavage LiP catalyzes the C a - C B cleavage of both B- l and B-O-4 dimeric model compounds2 6-2 7 , 4 2 , 4 4 , 6 1 , 6 2 as shown in Figures 1.6 and 1.7 respectively. Substituted phenylglycols are also cleaved at the C a - C B bond to give aldehydes.27 The C a - C B cleavage reaction of other model compounds catalyzed by L iP has not been reported so far. It is believed63 that C a - C B cleavage is probably the most important reaction in lignin biodegradation. It has also been reported,64, however, that lignin depolymerization does not occur to any significant extent even after significant oxidation. L iP was reported26 to be able to degrade lignin to give 4.5% of veratraldehyde, a product resulting from C a - C B cleavage. This result, however, does not indicate extensive C-C bond cleavage as the lignin used was methylated at the free hydroxy groups, which changed the properties of the lignin sample. 16 Figure 1.6 The mechanism of C 0-Cp cleavage of (3-1 model compounds: Figure 1.7 The mechanism of C a - C B cleavage of B-O-4 model compounds1 B. B - O - 4 bond cleavage and dealkylation reactions I n a d d i t i o n t o t h e C a - C B c l e a v a g e d i s c u s s e d a b o v e , L i P a l s o c a t a l y z e s t h e B - O - 4 e t h e r b o n d c l e a v a g e of B - O - 4 m o d e l c o m p o u n d s . T h e c l e a v a g e i s n o t a s i m p l e h y d r o l y s i s b u t a r e s u l t o f a s i n g l e e l e c t r o n o x i d a t i o n 5 9 , 6 5 a s s h o w n i n F i g u r e 1.8. A t t a c k of t h e C a - o r C y - h y d r o x y o n t h e B - r i n g c a t i o n r a d i c a l g i v e s c y c l i c i n t e r m e d i a t e s , h y d r o l y s i s o f w h i c h g i v e s t h e B - O - 4 e t h e r c l e a v a g e p r o d u c t s . L a t e r e x p e r i m e n t s s h o w e d t h a t r e a r r a n g e m e n t r e s u l t i n g f r o m o n e - e l e c t r o n o x i d a t i o n o c c u r s 6 6 , 6 7 a n d c y c l o h e x a d i e n e k e t a l s w e r e f o u n d i n t h e o x i d a t i o n o f l i g n i n m o d e l c o m p o u n d s b y L i P . 6 1 , 6 8 D e m e t h o x y l a t i o n i s a n i m p o r t a n t r e a c t i o n i n t h e m e t a b o l i s m o f l i g n i n b y P. Chrysosporium. A m e c h a n i s m f o r d e m e t h o x y l a t i o n w a s p r o p o s e d 5 9 b a s e d o n t h e r e s u l t s o f o x i d a t i o n o f a l k o x y b e n z e n e s b y L i P a s s h o w n i n F i g u r e 1.9. D e m e t h o x y l a t i o n h a s b e e n f o u n d t o o c c u r f o r a l k o x y b e n z e n e s 5 9 a n d f o r a C a - c a r b o n y l B - O - 4 d i m e r . 6 1 S m a l l a m o u n t s o f a d e m e t h o x y l a t i o n p r o d u c t w a s a l s o f o u n d i n t h e o x i d a t i o n o f v e r a t r y l a l c o h o l . 6 9 F r o m t h e e x p e r i m e n t a l e v i d e n c e a v a i l a b l e i t w o u l d a p p e a r t h a t d e m e t h o x y l a t i o n i s n o t a m a j o r r e a c t i o n o f l i g n i n p e r o x i d a s e c a t a l y s i s . I t b e c o m e s m o r e p r o n o u n c e d w h e n o t h e r , u s u a l l y m o r e f a v o u r a b l e , r e a c t i o n s a r e i m p o s s i b l e . C . Aromatic ring cleavage reactions T h e b i o l o g i c a l d e g r a d a t i o n o f l i g n i n i s h i g h l y o x i d a t i v e . 7 0 C h e m i c a l a n a l y s i s o f d e c a y e d l i g n i n s h o w e d 7 1 a d e c r e a s e i n t h e m e t h o x y l c o n t e n t a n d a n i n c r e a s e i n t h e c a r b o x y l c o n t e n t , i n d i c a t i n g o x i d a t i v e a r o m a t i c r i n g c l e a v a g e . D e g r a d a t i o n p r o d u c t s c o n t a i n i n g r e s i d u e s r e s u l t i n g f r o m a r o m a t i c r i n g o p e n i n g h a v e b e e n i s o l a t e d 7 2 " 7 4 a n d t h e r e i s e v i d e n c e 7 1 , 6 4 , 7 5 " 7 7 t h a t a r o m a t i c r i n g s c a n a l s o b e c l e a v e d w h i l e s t i l l b o u n d t o t h e l i g n i n 19 Figure. 1.8 The mechanism of B-O-4 cleavage of B-O-4 model compounds Figure 1.9 Possible mechanism of demethoxylation of lignin model compounds1 polymer. The molecular weight of k'gnin can remain virtually unchanged after extensive oxidation,64 suggesting that aromatic ring cleavage might be a critical step for lignin degradation. It is generally believed 6 3 , 7 1 , 7 8 , 7 9 that the aromatic rings of lignin are first demethylated and/or hydroxylated to give o-diphenols, cleavage of which by dioxygenases gives aliphatic carboxylic acids. Aromatic ring cleavage of a B-O-4 dimeric model compound by P. chrysosporium was reported.70 It has been found that aromatic rings of lignin model compounds can be cleaved by lignin peroxidase without the necessity of pre-demethoxylation or hydroxylation. L iP catalyzed aromatic ring cleavage of veratryl alcohol 6 9 , 8 0^ 2 and B-O-4 model compounds83"89 have been studied extensively and a mechanism was proposed based on isotope labelling experiments.86,89 As shown in Figure 1.10 using veratryl alcohol as an example, the ring opening is initiated by an one-electron oxidation. Coupling of molecular oxygen with the carbon-centered radical gives a peroxyl radical, further reaction of which gives the ring cleaved products. The isolation of direct ring-opening product (a muconate) from a B-O-4 dimer88 supports the mechanism described above. D. Benzylic alcohol oxidation Benzylic alcohol oxidation (Ca-oxidation) has been found as a reaction in the oxidation of lignin model compounds by LiP. Veratryl alcohol is oxidized to give veratraldehyde as a major product. Small amounts of C a-carbonyl products were also found from the oxidation of the C a-alcohol of dimeric lignin model compounds.61,83,90 (Figure 1.11) This reaction seems to be unfavourable for lignin model compound degradation since the aromatic ring is deactivated by the C a-carbonyl group for further oxidation. 22 ure 1.10 Proposed mechanism of aromatic ring cleavage of veratyl alcohol' Figure 1.11 Possible mechanism of Ca-oxidation of lignin model compounds Veratraldehyde is not oxidized by LiP. On the contrary, externally added veratraldehyde is rapidly reduced back to veratryl alcohol by the P. chrysosporium culture. The resulting veratryl alcohol is then metabolized to finally give carbon dioxide and water through aromatic ring cleavage and quinone intermediates.91 The role of the oxidation of veratryl alcohol, if it happens in vitro, is still not clear. Studies71 on white-rot decayed lignin showed that Ca-carbonyl groups were prominent. Though the A-ring is deactivated by the Ca-carbonyl (Figure 1.11), the B-ring is not affected and 0-C 4 (B-O-4 ether bond) cleavage resulting from B-ring oxidation is favoured in Ca-carbonyl model compounds.61 The aromatic ring opening of B-l and B-O-4 dimers is probably also facilitated by Ca-oxidation. Another possible consequence of Ca-oxidation is the cleavage of Ca-hemicellulose ether bonds and other Ca-ether bonds in lignin.91 The cleavage of Ca-ether bonds has been demonstrated for model compounds.61 Shimada92 proposed that C0-oxidation was important for lignin biodegradation since it could destroy the chiral centers at C a and Cg thus reducing the complexity of lignin. It was shown that Ca-oxidation of lignin enhances its depolymerization. The explanation was that the decreased steric complexity following Ca-oxidation opens up the polymer, making it more accessible to Ugnin degrading enzymes. Recent work36,93 showed that lignin model compound degradation was not stereospecific. The proposed mechanism of lignin degradation by LiP also suggests that the stereocomplexity of lignin does not hinder its degradation. E. Glycol formation from C a - C B double bonds C a - C B double bonds are found in the terminal chains of Ugnin. LiP can oxidize 25 OCH 3 OEt OEt Scheme 1.2 C a - C 6 double bond hydroxylation. these double bonds of lignin model compounds27-38,94 to give diols which are then further oxidized (Scheme 1.2). Isotopic labelling experiments61 showed that the oxygen of the CB-hydroxy of the resulting diol came from molecular oxygen. Diol formation from C a - C 6 double bonds were also found in the metabolism of lignin model compounds by P. chrysosporium cultures.95'98 The formation of diols via epoxides is a well known reaction of cytochromes P-450 which are monooxygenases and function by molecular oxygen activation.99 It seems unlikely that diol formation described above follows the mechanism of P-450 catalysis. In order for oxygen atom transfer to occur, and for epoxide to be formed, the substrate must closely approach the Fe™=0 bond of the protohemin at the active site of the heme protein. Due to the complexicity of lignin structures, such steric requirements cannot be met during lignin oxidation. F. Hydroxylation of benzylic methylene groups Lignin peroxidase can catalyze the hydroxylation of benzylic methylene groups (C a-hydroxylation). 3 6 , 3 8 , 4 5 1 8 0 from 1 8 0 2 was found to be incorporated into the C a -hydroxy group of the product under aerobic conditions, while under anaerobic 26 conditions, the C a-hydroxy oxygen was from water, suggesting a cation radical mechanism as shown in Figure 1.12. Ca-hydroxylation of model compounds was also found to be catalyzed by P. chrysosporium cultures.100,101 The importance of this reaction in lignin degradation is still not clear. G. Degradation and/or oxidation of lignin polymers Tien and Kirk first studied the degradation of lignin by L i P . 2 6 Spruce lignin and milled wood lignin (MWL) of birch were methylated at the free hydroxyl groups with 1 4 CH 3 I . Veratraldehyde (4.5%) was isolated from spruce lignin, 0.6% veratraldehyde and 0.4% syringaldehyde were isolated from birch lignin after incubation with lignin peroxidase. These results, however, do not provide convincing evidence that lignin peroxidase alone can depolymerize lignin since fundamental properties of the lignin were changed by methylation. The absence of free hydroxy groups makes the repolymerization more difficult. It is apparent that the methylated precursors of lignin biosynthesis (Scheme 1.1, 1, 2, and 3) cannot be polymerized to give methylated lignin or similar products under the conditions of lignin biosynthesis. Polymerization of lignin by enzymes from P. chrysosporium has been reported.102 Glenn et aP1 also reported the degradation of lignin by L iP in D M F . It is not clear if lignin depolymerization occurs in aqueous solution. Dordick et al103 reported that HRP and other peroxidases are able to vigorously depolymerize both natural and synthetic lignins in organic solvent, but are unable to do so in aqueous solution. A reassessment which questioned Dordick's experiment was reported later.104 Treatment of two synthetic lignins and one 1 4 C -methylated spruce lignin with crude lignin peroxidase105 decreased their degradation by Xanthomonas sp strain 99. Both polymerization and depolymerization was found for 28 the 14C-methylated spruce lignin while only polymerization was found for the synthetic lignins. Lignin peroxidase is unable to depolymerize lignin completely for as yet unknown reasons. Mechanism(s) must exist which can prevent the repolymerization of lignin by L iP or other enzymes. Though the degradation of lignin by P. chrysosporium is highly oxidative, there are enzymes in P. chrysosporium cultures which accomplish the reduction of acids, aldehydes, and quinones. The importance of these reductive reactions has been discussed recentiy.106 1.6 Biomimetic approaches in lignin biodegradation Lignin peroxidase is a non-specific enzyme containing protohemin as its prosthetic group. The hydrogen peroxide-oxidized intermediate, oxoiron(IV) protoporphyrin cation radical, is a powerful oxidizing species which can initiate one-electron oxidation of its substrates. Due to the heterogeneous structure of lignin, it seems unlikely that L iP will have an "active site" which can bind all the substructures of lignin and catalyze oxidations at its active site. Metalloporphyrins, therefore, could be good models for lignin peroxidase. These biomimetic catalysts may have advantages to the enzymes. Natural lignin is a three dimensional, water-insoluble high molecular weight polymer. Even the modified, water soluble Ugnin (such as Ugnosulfate) have compact, spheric structures in solution. The exponent a in the Mark-Houwink equation is about 0.1-0.5, corresponding to an intermediate form between an Einstein sphere and a compact coil. So a large part of lignin is not exposed to the active site of an enzyme during lignin degradation. Interaction of the Ugnin polymer and lignin degrading 29 enzymes is difficult to visualize. Being small molecules, metalloporphyrins, as lignin peroxidase models, may have an advantage in that their interactions with the lignin substrate may be easier than that with the enzyme. The properties of the metalloporphyrins, such as redox potential and solubility in organic or aqueous solvents, can be changed as desired by structural modification. Most metaUoporphyrins can be obtained in large quantities relatively easily. In contrast, lignin peroxidase is expensive and difficult to obtain in large quantity. Finally, enzymes are usually rather labile and very sensitive to pH, temperature, and solvent. Metalloporphyrins, on the other hand, can be very stable and can be used under extreme conditions. As lignin peroxidase catalyzes reaction by one-electron oxidation, electrochemical methods, which can initiate one-electron oxidation at the electrode, can also be used for biomimetic approaches to lignin biodegradation. 1.6.1 Degradation of lignin model compounds by iron porphyrins v Shimada et aP° first studied the degradation of a 6-1 model compound, l,2-bis(4-ethoxy-3-methoxyphenyl)propane-l,3-diol by iron tetraphenylporphyrin chloride (10, TPPFeCl, Figure 1.13) in the presence of t-butyl hydroperoxide (t-BuOOH) or iodosylbenzene in methylene chloride. The reaction was similar to that catalyzed by lignin peroxidase. Experiments107 in the presence of 1 8 0 2 or lyo showed the same isotopic incorporation pattern as that of the enzyme catalyzed reaction, indicating similar mechanisms for the two reactions. The degradation of a B-l model compound by protohemin (11, Figure 1.13) and hydrogen peroxide was also studied,108 C a-C 6 cleavage and Ca-oxidation were found to be the major reactions. Protohernin was also 30 12 M=Fe 14 M=Fe 13 M=Mn 15 M=Mn Figure 1.13 The structure of some metalloporphyrins shown to be able to catalyze the aromatic ring cleavage of veratryl alcohol.81 Shirnada et al109 studied the degradation of B-l, B-O-4, and B-5 dimers by piotohemin. The cation radical mechanism for the Ca-Cg cleavage of the B-l dimer was found to be supported by kinetic studies. While the B-l dimer was oxidatively cleaved by protohemin in a manner similar to the enzyme catalyzed reaction, the degradation of the B-O-4 dimer, 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether, gave products different from the enzyme catalyzed reaction.27 Wood et a/ 1 1 0 also reported the oxidation of a Ca-carbonyl B-O-4 model compound by a number of iron porphyrins, mcluding TPPFeCl, iron m&S0-tetra(2-methylphenyl)porphyrin, iron me$t?-temi(4-sulfonatophenyl)porphyrin, iron me5t9-tetra(4-N,N,N-trimemylanilinium)poiphyrin, and iron me&>tetra(N-memyl-4-pyridyl)porphyrin. 1.6.2 Oxidation of lignin model compounds by inorganic one-electron oxidants It was reported43,60 that eerie ammonium nitrate and tris(phenanthrolme)iron(III) could oxidize l,2-bis(3,4-d^ethoxyphenyl)propane-l,3-diol to give veratraldehyde. An initial one-electron oxidation was suggested as the mechanism of the reaction. Further studies by Fisher and Dershem111 showed that eerie ammonium nitrate did not cleave the B-l model compound by one-electron oxidations. Instead, coordination of the a-hydroxy group of the B-l dimer to cerium and decomposition of the resulting complex direcdy gave the product. Huynh112 reported the oxidation of several model compounds by copper peroxydisulfate in 5:1 acetic acid/water at reflux temperature. Veratrylglycerol-B-guaiacyl ether can be degraded by this system giving products similar to that of the LiP catalyzed reaction. In the degradation of dihydroanisoin (l,2-di(4-methoxyphenyl)-l,2-32 ethanediol) and veratryl alcohol, side reactions occurred. Veratrylglycerol-B-guaiacyl ether was reported113 to be cleaved at the C a -C B bond by Co(H)/Mn(H) in 80% acetic acid under oxygen (4% oxygen in nitrogen, 500 psi) at 170°C. The Ca-Cg cleavage resulted from an initial single-electron oxidation. Treating the B-l model compounds, dihydroanisoin and l^-di(4-methoxyphenyl)propane-l,3-diol, under the same condition gave only acid-catalyzed dehydration products. 1.6.3 Electrochemical oxidation of lignin model compounds Electrochemical methods are useful tools for studying reactions involving cation radical intermediates. It is known114 that 3,5-dimethoxy-4-hydroxy-a-methylbenzyl alcohol can be oxidized by HRP in the presence of excess hydrogen peroxide to give 3,5-dimethoxy-p-benzoquinone. Phenoxy radicals of 3,5-dimethoxy-4-hydroxy-a-methylbenzyl alcohol and of 3,5-dimethoxy-4-hydroxyacetophenone were detected during the reaction. When less than one equivalent of hydrogen peroxide was used the phenoxy radical of 3,5-dimethoxyacetophenone was not detected but 3,5-dimethoxy-p-benzoquinone was still found as a product. By oxidizing the substrate at different potentials, electrochemical techniques showed that the quinone can be formed directly from the starting material without requiring 3,5-dimethoxy-4-hydroxyacetophenone as an intermediate.115 The demethylation mechanism of 3,5-dimethoxy-4-hydroxyacetophenone was also readily established by electrochemical methods.116 Electrochemical oxidation of alkoxyphenols1"*119 indicated that two pathways were involved. They can be oxidized by removing two electrons to give the phenoxonium ions, reaction of which with water or other nucleophiles gives the final products. The second pathway, which is favoured in the presence of base, involves one-electron 33 oxidation to give a phenoxy radical, subsequent coupling of the radical gives polymerized products. Limosin et al120 have studied the electrochemical oxidation of over 20 dimeric lignin model compounds in acetonitrile. The major reaction was polymerization. The oxidation of a B-O-4 model compound in aqueous acetonitrile,121 however, gave cleavage products. The electrochemical oxidation of lignin in basic aqueous solution has been studied.122 A decrease in phenyl groups and an increase in carboxylic acid group content was found and aromatic ring cleavage was also found. A review addressing the practical aspects of the electrochemistry of lignin was prepared by Chum et al.123 Lignins can be effectively degraded to small molecules by anodic oxidation. The reactions are probably different from LiP catalyzed reactions as the reaction conditions (pH) are different. 1.7 The purpose of this study Although simple metalloporphyrins have been used as models for lignin peroxidase,81,91,107"110 metalloporphyrin catalyzed degradation of model compounds representing all the major substructures of lignin have not been reported. Protohemin81 (11) and iron mew-tetraphenylporphyrin chloride (TPPFeCl, 10) were able to mimic lignin peroxidase in the degradation of veratryl alcohol and B- l model compounds. The degradation of a B-O-4 model compound, which represents the most abundant substructure of lignin, by protohemin and lignin peroxidase gave different products.109 The oxidation of this important model compound by metalloporphyrins needed to be re-examined. 34 The degradation of B-5 and 5-5' biphenyl compounds by iron porphyrins have not been studied. Reactions such as hydroxylation of benzylic methylene groups and glycol formation from C a - C 6 double bonds have not been reported for metalloporphyrin catalyzed reactions. These reactions have to be studied in order to demonstrate that metalloporphyrins mimic lignin peroxidase in all cases. The metalloporphyrins used in previous studies81,90,107"110 suffer a great disadvantage in that they are very unstable towards excess oxidants. A catalyst to substrate ratio of 1:10 had to be used.109 In addition, most of the reactions were performed in organic solvents and reactions in organic and aqueous solutions may follow different pathways. Dramatic solvent effect has been observed for enzyme catalyzed oxidations of lignin. 1 0 3 Biotechnology holds great potential in the pulp and paper industry. Biomimetic catalysts such as metalloporphyrins provide an alternative for using enzymes and microorganism cultures in pulping and pulp bleaching processes. The metalloporphyrins, however, have to be made stable and water soluble before they can be used in industry. Water soluble, sterically protected metalloporphyrins, the iron(ILT) and manganese(III) complexes of me5o-tetra(2,6-dichloro-3-sulfonatophenyl)porphyrin (TDCSPPFeCl, 12; TDCSPPMnCl, 13; Figure 1.13), and meso-tetta(2,6-dichloro-3-sulfonatophenyl)-fJ-octachloroporphyrin (Cl 1 6TSPPFeCl, 14; Cl 1 6 TSPPMnCl, 15; Figure 1.13) have been synthesized and used as models for lignin peroxidase. Factors affecting the catalytic activities of these metalloporphyrins have been studied. The degradation of a number of lignin model compounds, including veratryl alcohol, B - l , B-0-4, B-5, 5-5' biphenyl, phenylpropane and phenylpropene model compounds, are presented. Preliminary studies on the oxidation of environmental pollutants, pyrene and 35 2,4,6-trichlorophenol, are reported. Metallophthalocyanines, a class of compounds closely related to the metalloporphyrins, have also been examined as models for lignin peroxidase. Manganese(H)-complexes and veratryl alcohol have been suggested to function as mediators for the manganese-dependent peroxidase and Ugnin peroxidase, respectively. Metalloporphyrins and electrochemical techniques have been used to study the mediating effects of manganese(II)-complexes and veratryl alcohol. 36 References for Chapter 1 1. T. Higuchi, M . Shimada, F. Nakatsubo, and M . Tanahashi, Wood Sci. Technol. 11, 153, 1977. 2. E. Adler, Wood Sci. Technol. 11, 169, 1977. 3. H . Nimz, Angew. Chem. 86, 336, 1974. 4. D. W. Ribbons, Phil. Trans. R. Soc. Lond. A321, 485, 1987. 5. D. Feldman, L . Lacasse, and M . Beznaczuk, Prog. Polym. Sci. 12, 271, 1986. 6. B. E. Dale, Trends in Biotechnol. 5, 287, 1987. 7. M . Tanaka, R. Matsuno, Enzyme Microb. Technol. 7, 197, 1985. 8. G. F. Leatham, G. C. Myers, T. H . Wegner, and R. A . Blanchette, In proceedings of the Fourth International Conference on Biotechnology in the Pulp and Paper Industry. Raleigh, N.C., USA, 1989, in press. 9. M . G. Paice, L . Jurasek, C. Ho, R. Bourbonnais, and F. Archibald, see Ref. 8. 10. P. S. Skerker, R. L . Farrell, D. Dolphin, F. Cui and T. Wijesekera, see Ref. 8. 11. D. L . Crawford, and R. L . Crawford, Enzyme Microb. Technol. 2, 11, 1980. 12. R. L . Crawford, and D. L . Crawford, Enzyme Microb. Technol. 6, 434, 1984. 13. R. Vicuna, Enzyme Microb. Technol. 10, 646, 1988. 14. J. Trojanowski, K , Haider, and V . Sundman, Arch. Microbiol. 114, 149, 1977. 15. T. K . Kirk, and E. Adler, Acta Chem. Scand. 24, 3379, 1970. 16. T. K . Kirk, Holzforschung, 29, 99, 1975. 17. T. K . Kirk, W. J. Connors, R. D. Bleam, W. F. Hackett, J. K . Zeikus, Proc. Natl. Acad. Sci. USA, 72, 2515, 1975. 37 18. T. K . Kirk, and R. L . Farrell, Ann. Rev. Microbiol. 41, 465, 1987. 19. B. D. Faison, T. K . Kirk, and R. L . Farrell, Appl. Environ. Microbiol. 52, 251, 1986. 20. M . S. A . Leisola, D. C. Ulmer, R. Waldner, A . Fiechter, J. Biotechnol. 1, 331, 1984. 21. B. D. Faison, and T. K . Kirk, Appl. Environ. Microbiol. 49, 299, 1985. 22. D. C. Ulmer, M . S. A . Leisola, and A . Fiechter, J. Biotechnol. 1, 13, 1984. 23. P. Hall, Enzyme Microb. Technol. 2, 170, 1980. 24. T. K . Kirk, M . D. Muzuch, and M . Tien, Biochem. J. 266, 455, 1985. 25. T. K . Kirk, F. Nakatsubo, and I. D. Raid, Biochem. Biophys. Res. Comm. I l l , 200, 1983. 26. M . Tien, and T. K . Kirk, Science, 221, 661, 1983. 27. J. K . Glenn, M . A . Morgan, M . B. Mayfield, M . Kuwahara, and M . H . Gold, Biochem. Biophys. Res. Commun. 114, 1077, 1983. 28. M . Kuwahara, J. K . Glenn, M . A. Morgan, and M . H . Gold, FEBS Letters, 169, 247, 1984. 29. J. K . Glenn, and M . H . Gold, Arch. Biochem. Biophys. 242, 329, 1985. 30. M . S. A . Leisola, B. Kozulic, F. Menssdoerffer, and A . Fiechter, J. Biol. Chem. 262, 419, 1987. 31. R. L . Farrell, K . E. Murtagh, M . Tien, M . D. Mozuch, and T. K. Kirk, Enzyme Microb. Technol. 11, 322, 1989. 32. F. Tonon, and E. Odier, Appl. Environ. Microbiol. 54, 466, 1988. 33. M . Tien, see Ref. 8. 34. H . Wariishi, K . Valli , and M . H . Gold, Biochem. 28, 6017, 1989. 35. M . H . Gold, H . Wariishi, K . Vall i , M . B. Mayfield, V . J. Nipper, and D. 38 Pribnow, see Ref. 8. 36. M . Tien, and T. K . Kirk, Proc. Nad. Acad. Sci. USA, 81, 2280, 1984. 37. J. Trailer, J. D. G. Smit, M . S. A . Leisola, J. Kallen, K . H . Winterhalter, and A. Fiechter, Bio/technol. 6, 571, 1988. 38. V . Renganathan, K . Mik i , and M . H . Gold, Arch. Biochem. Biophys. 241, 304, 1985. 39. V . Renganathan, and M . H . Gold, Biochem. 25, 1626, 1986. 40. D. Kuila, M . Tien, J. A . Fee, and M . R. Ondrias, Biochem. 24, 3394, 1985. 41. L . A . Anderson, V . Renganathan, T. M . Loehr, and M . H . Gold, Biochem. 26, 2258, 1987. 42. L . A . Anderson, V . Renganathan, A . A . Chiu, T. M . Loehr, and M . H . Gold, J. Biol. Chem. 260, 6080, 1985. 43. P. J. Harvey, H . E. Schoemaker, R. M . Bowen, and J. M . Palmer, FEBS Letters, 183, 13, 1985. 44. K . E. Hammel, M . Tien, B. Kalyanaraman, and T. K . Kirk, J. Biol. Chem. 260, 8348, 1985. 45. V . Renganathan, K . Mik i , and M . H . Gold, Arch. Biochem. Biophys. 246, 155, 1986. 46. J. M . Palmer, P. J. Harvey, and H . E. Schoemaker, Phil. Trans. R. Soc. Lond. A321, 495, 1987. 47. M . Tien, T. K . Kirk, C. Bull, and J. A . Fee, J. Biol. Chem. 261, 1687, 1986. 48. A . Pasczynski, V . -B . Huynh, and R. Crawford, Arch. Biochem. Biophys. 244, 750, 1986. 49. Y . Mino, H . Wariishi, N . J. Blackburn, T. M . Loehr, and M . H . Gold, J. Biol . Chem. 263, 7029, 1988. 39 50. H . Wariishi, L . Akileswaran, and M . H . Gold, Biochem. 27, 5365, 1988. 51. A . Haars, and A . Huttermann, Naturwissenschaften, 67, 39, 1980. 52. A . Huttermann, C. Herche, and A . Haars, Holzforschung, 34, 64, 1980. 53. A . Haars, and A- Huttermann, Arch. Microbiol. 125, 233, 1980. 54. N . Morohoshi, H . Wariishi, C. Muraiso, T. Nagai, and F. Haraguchi, Mokuzai Gakkaishi, 33, 218, 1987. 55. N . Morohoshi, and T. Haraguchi, Mokuzai Gakkaishi, 33, 143, 1987. 56. N . Morohoshi, K . Fujita, H . Wariishi, and H . Haraguchi, Mokuzai Gakkaishi, 33, 310, 1987. 57. S. Kawai, T. Umezawa, M . Shimada, T. Higuchi, K . Koide, T. Nishida, N . Morohoshi, and T. Haraguchi, Mokuzai Gakkaishi, 33, 792, 1987. 58. H . Wariishi, N . Morohoshi, and T. Haraguchi, Mokuzai Gakkaishi, 33, 892, 1987. 59. P. J. Kersten, M . Tien, B . Kalyanaraman, and T. K . Kirk, J. Biol. Chem. 260, 2609, 1985. 60. H . E. Schoemaker, P. J. Harvey, R. M . Bowen, and J. M . Palmer, FEBS Letters, 183, 7, 1985. 61. T. K . Kirk, M . Tien, P. J. Kersten, M . D. Mozuch, and B. Kalyanaraman, Biochem. J. 236, 279, 1986. 62. K . E . Hammel, B. Kalyanaraman, and T. K . Kirk, Proc. Natl. Acad. Sci. USA, 83, 3708, 1986. 63. C.-L. Chen, and H . - M . Chang, in Biosynthesis and Biodegradation of Wood Components, Ed. T. Higuchi, San Diego, C A : Academic Press, 1985, Chp. 19. 64. T. K . Kirk, and H . - M . Chang, Holzforschung, 28, 217, 1974. 65. T. Umezawa, and T. Higuchi, Agric. Biol. Chem. 48, 1917, 1984. 40 66. K. Mik i , V . Renganathan, and M . H . Gold, FEBS Letters, 203, 235, 1986. 67. V . - B . Huynh, A . Paszczynski, P. Olson, and R. Crawford, Arch. Biochem. Biophys. 250, 186, 1986. 68. S. Kawai, T. Umezawa, and T. Higuchi, FEBS Letters, 210, 61, 1987. 69. S. D. HaemmerU, H . E. Schoemaker, H . W. H . Schmidt, and M . S. A . Leisola, FEBS Letters, 220, 149, 1987. 70. T. Umezawa, and T. Higuchi, FEBS Letters, 182, 257, 1985 . 71. T. K . Kirk, and H . - M . Chang, Holzforschung, 29, 56, 1975. 72. C.-L. Chen, H . - M . Chang, and T. K . Kirk, Holzforschung, 36, 3, 1982. 73. C.-L. Chen, H . - M . Chang, and T. K . Kirk, J. Wood Chem. Technol. 3, 35, 1983. 74. D. Tai, M . Terasawa, C.-L. Chen, H . - M . Chang, and T. K . Kirk, in Recent advances in Lignin Biodegradation Research, Ed. T. Higuchi, H . - M . Chang, and T. K. Kirk, 1983, p44. 75. M . G. S. Chua, C.-L. Chen, H . - M . Chang, and T. K . Kirk, Holzforschung, 36, 165, 1982. 76. K . Haider, H . W. Kern, and L . Ernst, Holzforschung, 39, 23, 1985. 77. H . Junshekar, C. Brown, T. Haltmeier, M . S. A . Leisola, and A . Fiechter, Arch. Microbiol. 132, 14, 1982. 78. R. L . Crawford, Lignin Biodegradation and Transformation, John Wiley & sons, New York, 1981, p42, pl07. 79. D. L . Crawford and R. L . Crawford, Enzyme Microb. Technol. 2, 11, 1980. 80. M . S. A . Leisola, B . Schmidt, V . Thanei-Wyss, and A . Fiechter, FEBS Letters, 189, 267, 1985. 81. M . Shimada, T. Hattori, T. Umezawa, T. Higuchi, and T. Okamoto, Colloq. INRA 40 ( Lignin Enzymic Microb. Degrad. ), 151, 1987. 41 82. M . S. A . Leisola, S. D. Haemmerli, J. D. G. Smit, J. Trailer, R. Waldner, H . E. Schoemaker, and H . Schmidt, Collq. INRA 40 ( Lignin Enzymic Microb. Degrad. ), 81, 1987. 83. T. Umezawa, M . Shimada, T. Higuchi, and K . Kusai, FEBS Letters, 205, 287, 1986. 84. T. Umezawa, and T. Higuchi, FEBS Letters, 205, 293, 1986. 85. K . Mik i , V . Renganathan, M . B . Mayfield, and M . H . Gold, FEBS Letters, 210, 199, 1987. 86. T. Umezawa, and T. Higuchi, Colloq. INRA 40 ( Lignin Enzymic Microb. Degrad. ), 63, 1987. 87. K. Mik i , V . Renganathan, M . B . Mayfield, and M . H . Gold, Colloq. INRA 40 (Lignin Enzymic Microb. Degrad. ), 69, 1987. 88. T. Umezawa, and T. Higuchi, Agric. Biol. Chem. 51, 2281, 1987. 89. K. Mik i , R. Kondo, V . Renganathan, M . B. Mayfield, and M . H . Gold, Biochem. 27, 4787, 1988. 90. M . Shimada, T. Habe, T. Umezawa, and T. Higuchi, Biochem. Biophys. Res. Commun. 122, 1247, 1984. 91. M . S. A . Leisola, S. D. Haemmerli, R. Waldner, H . H . Schoemaker, H . W. H . Schmidt, and A . Fiechter, Cellulo. Chem. Technol. 22, 267, 1988. 92. M . Shimada, in Lignin Biodegradation: Microbiol. Chem. and Potential Appl. Ed. T. K . Kirk, T. Higuchi, and H . - M . Chang, C R C Press, 1980, ppl95. 93. F. Nakatsubo, I. D. Reid, and T. K . Kirk, Biochim. Biophys. Acta, 719, 284, 1982. 94. M . H . Gold, M . Kuwahara, A . A . Chiu, .• and J. K . Glenn, Arch. Biochem. Biophys. 234, 353, 1984. 42 95. T. Umezawa, F. Nakatsubo, and T. Higuchi, Agric. Biol . Chem. 47, 2677, 1983. 96. T. Higuchi, Wood Res. 67, 47, 1981 . 97. T. Katagama, and T. Higuchi, Abst. 25th Symp. Lignin Chem. Fukuoko, Japan, 1980, pp5. 98. F. Nakatsubo, T. K . Kirk, M . Shimada, and T. Higuchi, Arch. Microbiol. 128, 416, 1981. 99. B. W. Griffin, J. A . Peterson, and R. W. Estabrook, in The Porphyrins, Ed. D. Dolphin, Academic Press, 1979, Vol . VII, p333. 100. G. P. Goldsby, A . Enoki, and M . H . Gold, Arch. Microbiol. 128, 190, 1980. 101. A . Enoki, and M . H . Gold, Arch. Microbiol. 132, 123, 1982. 102. S. D. Haemmerli, M . S. A . Leisola, and A . Fiechter, FEBS Microbiol. Letters, 35, 33, 1986. 103. J. S. Dordick, M . A . Marietta, and A . M . Klibanov, Proc. Natl. Acad. Sci. USA, 83, 6255, 1986. 104. N . G. Lewis, R. A . Razal, and E. Yamamoto, Proc. Natl. Acad. USA, 84, 7925, 1987. 105. H . W. Kern, and T. K. Kirk, Appl. Environ. Microbiol. 53, 2242, 1987. 106. H . E. Schoemaker, E. M . Meijer, M . S. A . Leisola, S. D. Haemmerli, R. Waldner, D. Sanglard, and H . W. H . Schmidt, in Plant Cell Wall Polymers, Biogenesis and Biodegradation. Ed. N . G. Lewis, and M . G. Paice, Am. Chem. Soc. Washington DC, 1989, chp33. 107. T. Habe, M . Shimada, T. Okamoto, B . Panijpan, and T. Higuchi, J. Chem. Soc. Chem. Commun. 1323, 1985. 108. T. Habe, M . Shimada, and T. Higuchi, Mokuzai Gakkaishi, 31, 54, 1985. 109. M . Shimada, T. Habe, T. Higuchi, T. Okamoto, and B . Panijpan, Holzforschung, 43 41, 277, 1987. 110. J. W. Wood, B. R. Folsom, and W. W. Eudy, in Biotechnology in the Pulp and Paper Industry ( 3rd international conference), Stockholm, Sweden, June 16-19, 1986, p31. 111. T. M . Fisher, and S. M . Dershem, J. Org. Chem. 53, 1504, 1988. 112. V . - B . Huynh, Biochem. Biophys. Res. Commun. 139, 1104, 1986. 113. R. Dicosimo, and H.-C. Szabo, J. Org. Chem. 53, 1673, 1988. 114. M . Young, and C. Steelink, Phytochem. 12, 2851, 1973. 115. C. Steelink, and W. E. Brritton, Tetrahedron Letters, 33, 2869, 1974. 116. W. E. Britton, and C. Steelink, Tetrahedron Letters, 33, 2873, 1974. 117. F. Sundholm, Holzforschung, 36, 1982. 118. F. J. Vermillion, and J. A . Pearl, J. Electrochem. Soc. I l l , 1392, 1964. 119. B. Chaband, F. Sundholm, and G. Sundholm. Electrochimica Acta, 23, 659, 1978. 120. D. Limosin, G. Pierre, and G. Canquis, Holzforschung, 39, 91, 1985. 121. L . Y . Dudenko, M . I. Anisimova, V . V . Yanilxin, V . A . Babkin, S. Z. Ivanova, L . N. Spirridonova, and V . G. Gorokhova, Khim. Drev. 5, 70, 1985. 122. D. Limosin, G. Pierre, and G. Cauguis, Holzforschung, 40, 31, 1986. 123. H . L . Chum, D. W. Sopher, and H . Schroeder, in Fundamentals of Thermochemical Biomass Conversion, Ed. R. P. Overend, R. A . Milne, and L . K . Mudge, Elsevier Appl. Sci., New York, 1985, chp. 62. 44 Chapter 2 Metalloporphyrins as lignin peroxidase models 2.1 Results and discussion 2.1.1 Stability of the metalloporphyrins towards excess oxidant Enzymes are proteins that catalyze chemical reactions in biological systems with enormous catalytic power. Most enzymes are substrate specific and this substrate specificity is determined by the three-dimensional structure of the enzyme active site, which is provided by a certain conformation of the protein. The amino acid residues near the active site also provide a special chemical environment (/. e. hydrophobic, hydrophillic) to facilitate the reaction, and to contribute special groups participating in the catalytic process. Lignin peroxidase is an unusual enzyme which has litde or no substrate specificity. It works by one-electron oxidation and oxidizes a wide variety of substrates, including various Ugnin model compounds, alkoxybenzenes, and polycyclic aromatic hydrocarbons. It is unlikely that lignin peroxidase wiU have an "active site" which can bind all the Ugnin substructures and other substrates. The protein part of lignin peroxidase, therefore, may not be as important as that of other enzymes for the catalytic process. However, it probably plays an important role in protecting the 45 oxidized prosthetic group of the enzyme. Without such protection, simple iron porphyrins such as protohemin are rapidly self-destroyed in the presence of excess oxidants. Metalloporphyrins, especially iron, manganese, chromium, and ruthenium complexes of meso-tetraphenylporphyrin derivatives (TPP's) have been used as models for peroxidases and cytochrome P-450. Various substituents can be introduced at the phenyl ring and/or the B-position of TPP's to increase their stability. The metal complexes of the following porphyrins have been used as models for various enzymes: /neso-tettamesitylporphyrin, meso-tetta(pentafluorophenyl)porphyrin, meso-tetra(o-phenylphenyl)porphyrin, me5o-tetra(2,6-dichlorophenyl)porphyrin, m&sotetra(2,6-dichlorophenyl)-B-octachloroporphyrin, and me£0-tetra(2,6-dichlorophenyl)-B-octabromoporphyrin. Chloro-substituted, water soluble TPP derivatives, 12, 13, 14, and 15 Q?igufe 1.13) were used in this study. Iron me,St>tetra(4-sulfonatophenyl)porphyrin chloride (TSPPFeCl) was synthesized and used for comparison. Figure 2.1 shows the relative stability of TDCSPPFeCl (12), Cl : 6 TSPPFeCl (14), TDCSPPMnCl (13), Cl 1 6 TSPPMnCl (15) (Figure 1.13), and TSPPFeCl at pH 3 towards 100 fold excess (molar ratio) of hydrogen peroxide in the presence of veratryl alcohol. TSPPFeCl is the least stable and about 70% of it is destroyed in 10 minutes. The chloro-substituted metalloporphyrins 12-15 are much more stable and less than 10% of them are destroyed under the same condition. Figure 2.1 shows clearly that both manganese porphyrins are more stable than the iron porphyrins and the octachloroporphyrins (TDCSPPFeCl, TDCSPPMnCl) are more stable than their corresponding B-octachloro analogues (Cl 1 6TSPPFeCl, Cl 1 6 TSPPMnCl). This stability order seems to be in conflict with that reported in the literature1 and the reasons are not clear. Iron me^o-tetra(2,6-dichlorophenyl)-B-octabromoporphyrin was more stable than 46 A «> 1.00 -0.20 H 1 1 1 1 1 > 0 2 4 6 8 10 Time (min) Figure 2.1 The relative stability of metalloporphyrins (7.5x10* M) towards 100 fold excess of hydrogen peroxide (7.5x10"* M) at pH 3 (0.1 M phosphate buffer) in the presence of veratryl alcohol (2.5xl03 M) at room temperature: 1, TDCSPPMnCl; 2, Cl16TSPPMnCl; 3, TDCSPPFeCl; 4, Cl16TSPPFeCl; 5, TSPPFeCl; iron mejo-tetra(2,6-dichlorophenyl)porphyrin, iron meso-tetramesitylporphyrin, and iron mejo-tetra(pentafluorophenyl)porphyrin towards pentafluoroiodosylbenzene in organic solvents. The stability of the metalloporphyrins depends on the pH of the solvent and Figure 2.2 shows the relative stability of the metalloporphyrins at pH 10 in the presence of veratryl alcohol and 100 fold excess (molar ratio) of hydrogen peroxide. The results are plotted as ( A S o n ! l ) / ( A S o r J 0 against time t, where (A S o r e t ) t is the Soret absorption at time t, and ( A ^ Q is the Soret absorption at time 0. The stability order is the same as that at pH 3: TDCSPPMnCl > Cl 1 6 TSPPMnCl > TDCSPPFeCl > Cl 1 6 TSPPFeCl. It can be seen by comparing Figure 2.1 and Figure 2.2 that all the metalloporphyrins (12-15) are more stable at lower pH. It was reported2, however, that manganese porphyrins were more stable towards oxidant at higher pH in the absence of substrate. The stability of TDCSPPFeCl towards mCPBA in the absence of veratryl alcohol is shown in Figure 2.3. As expected, the iron porphyrin is indeed more stable at higher pH in the absence of substrates. When hydrogen peroxide is the oxidant, however, TDCSPPFeCl is relatively more stable at lower pH (Figure 2.4). The stability of the metalloporphyrins also depends on the type of oxidant used. As shown in Figure 2.5, Cl 1 6 TSPPFeCl is much more stable towards t-BuOOH than towards hydrogen peroxide. These results indicate that both the relative and absolute stabilities of the chloro-substituted porphyrins depend on several factors including the type of oxidant, pH, and the presence or absence of substrates. 48 ( A S o r e t ) t / ( A Soret) A 1.00 0.80-0.60-0.40 0.20-0.00 T(min.) Figure 2.2 The relative stability of the metalloporphyrins (7.5x10"* M) towards 100 fold excess of hydrogen peroxide (7.5x10^ M) at pH 10 (0.1 M phosphate buffer) in the presence of veratryl alcohol (2.5xl0' 3 M) at room temperature: 1, TDCSPPMnCl; 2, Cl 1 6 TSPPMnCl; 3, TDCSPPFeCl; 4, Cl . JSPPFeCl ; 49 ( Asowt)t/( Asoret)o A 1.00 J 0.0 1.0 2.0 3.0 4.0 5.0 Time (min.) Figure 2.3 The stability of TDCSPPFeCl ( l x l O 5 M) towards raCPBA ( l x l O 4 M) in 0.1 M phosphate buffer at room temperature in the absence of veratryl alcohol 50 (Asore t) t/( ASoret) 0 1.00 0.80 -0.60 -0.40 0.0 1.0 2.0 3.0 4.0 5.0 Time (min) Figure 2.4 The stability of TDCSPPFeCl (1x10 s M) towards hydrogen peroxide (1x10/ M) in 0.1 M phosphate buffer at room temperature in the absence of veratryl alcohol 51 (Asoret)t/(ASoret)0 A 1.00 0.90 0 . 8 0 -| 1 1 1 1 1 1 1 1 1 1 ^* 0.0 2.0 4.0 6.0 8.0 10.0 Time (min.) Figure 2.5 The stability of Cl 1 6 TSPPFeCl towards t-BuOOH and H 2 0 2 at pH 3 in the presence of veratryl alcohol at room temperature, the reaction mixture contains in 3 mL 0.1 M phosphate buffer 3x10"* mole of Cl 1 6 TSPPFeCl, 3X10"6 mole of t-BuOOH or H 2 0 2 , and lxlO" 5 mole of veratryl alcohol. 52 2.1.2 Oxidized intermediates of the metalloporphyrins Biological systems use a variety of heme containing enzymes to carry out various efficient and selective oxidations. Understanding the function and mechanism of these enzymes is of great theoretical and industrial importance. Synthetic metalloporphyrins have been extensively studied as biomimetic models for monooxygenases and peroxidases and have provided a means to isolate and characterize reactive intermediates implicated in these biological systems. Oxidation of iron porphyrins by iodosylbenzene or peracids at low temperature" gives an oxoiron(IV) porphyrin cation radical (corresponding to compound I of peroxidase), one electron reduction of which gives a second intermediate, oxoiron(IV) porphyrin (corresponding to compound H of peroxidase). Both intermediates are very reactive and are responsible for the oxidations catalyzed by heme.3* Two oxidized intermediates of manganese(III) porphyrins, oxomanganese(rV) and oxomanganese(V) were suggested to be involved in manganese(III) porphyrin catalyzed alkane hydroxylation and alkene epoxidation reactions. Oxomanganese(IV) porphyrins (or hydroxide adducts) seem to be fairly stable and have been characterized by UV-vis, resonance Raman, TR, and ESR spectroscopy.7"10 X-ray crystal structure of a manganese(rV) porphyrin p>oxo dimer, u-oxo bis[azidotetraphenylporphyrinato-manganese(IV)], has been reported.11 Manganese(iV) porphyrin-iodosylbenzene adducts, TPPMnIV-bis(iodosylbenzene) and a u.-oxo dimer, (TPPMn IV)2-0-bis(iodosylbenzene) have been shown to be able to functionalize both alkenes and alkanes.12 The second oxidized intermediate of manganese porphyrin, believed to be an oxomanganese(V) porphyrin, was suggested to be involved in manganese porphyrin catalyzed reactions.7,8 ,13 ,14 This species seems to be very unstable and has not been fully 53 characterized. When produced by reacting m&so-teti^ esitylporphyrin manganese hydroxide with /nCPBA at -78°C in methylene chloride, it decays to give the oxomanganese(IV) porphyrin in a few minutes.14 Physical and chemical evidence7,8,13,14 suggest that this second oxidized intermediate, which is believed to be an oxomanganese(V) porphyrin, is different from the well characterized oxornanganesefTV) porphyrin species. It is more reactive than oxomanganese(IV) porphyrins for alkene epoxidation. The stereoselectivity and isotope incorporation pattern of alkene epoxidation in the presence of lyo or 1802 catalyzed by oxomanganesetTV) porphyrin and the second oxidized species are different. OxomanganesetTV) porphyrins have ESR signals while the second intermediate is ESR silent It has been shown that decomposition of acylperoxomanganese(JJI)15,16 and manganese(IH)-peroxycarbonate porphyrins10 also gave oxomanganese(V) porphyrin species. The oxomanganese(V) porphyrin thus generated15 has two oxidizing equivalents over the manganese(III) porphyrin. All these facts supports the suggestion that the second oxidized intermediate of manganese porphyrin is an oxomanganese(V) complex. The oxidation of water soluble manganese porphyrins were also studied2,17', under basic conditions both manganese(IV) and oxomanganese(V) porphyrins were claimed to be produced. Oxomanganese(V) porphyrins seem to be more stable in basic aqueous solution than in organic solvents.2,17 Several oxidized intermediates of TDCSPPMnCl, Cl16TSPPMnCl, TDCSPPFeCl, and Cl16TSPPFeCl were observed by UV-vis spectroscopy. Due to the short life times and difficulties in handling these water soluble intermediates, no effort was made to characterize these intermediates by other physical methods. 54 2.1.2.1 Oxidized intermediates of TDCSPPMnCl A . Oxidation of TDCSPPMnCl by sodium hypochlorite TDCSPPMnCl was oxidized by sodium hypochlorite (NaCIO) at various pH's in the presence or absence of veratryl alcohol at room temperature to examine the stability and reactivity of the oxidized intermediates towards veratryl alcohol. The oxidation of TDCSPPMnCl was followed using a diode array UV-vis spectrometer. When NaCIO (lxlO- 6 mole) was added to TDCSPPMnCl (3X10"8 mole) in pH 2 phosphate buffer (3 mL), no spectral change was observed. Addition of another 5x10* mole of NaCIO to the solution caused a slow decrease of the 472 nm peak of TDCSPPMnCl and a new peak at 432 nm was slowly formed. The intensity of the 472 nm peak of TDCSPPMnCl decreased by half in about 15 minutes. At pH 3, the oxidation of TDCSPPMnCl by NaCIO to the new species (16) which had its Soret band at 432 nm was also slow and the 472 nm peak did not disappear completely in 30 minutes after NaCIO (5x10* mole) was added to TDCSPPMnCl (3x10"* mole). The oxidation is very fast at pH 7 and the addition of NaCIO (1x10* mole) to TDCSPPMnCl (3x10"* mole in 3 mL 0.1 M phosphate buffer) brought about an instant reaction giving 16. The intermediate 16 seems fairly stable and the 432 nm peak decreased very slowly at room temperature in air. Addition of veratryl alcohol (1x10* mole) to 16 caused its rapid reduction to give TDCSPPMnCl as shown in Figure 2.6. Veratryl alcohol was oxidized at the same time to give veratraldehyde, as seen by the increase of the absorption of veratraldehyde at 310 nm. The intermediate 16 was probably reduced by veratryl alcohol to give the Mn(III) porphyrin directly. Figure 2.6 shows clearly the presence of three isosbestic points at 55 Figure 2.6 UV-vis spectral changes when veratryl alcohol was added to the oxidized intermediate 16 (TDCSPPMn™^, 432 nm) in pH 7 phosphate buffer (0.1 M) at room temperature 390 nm, 456 nm, and 508 nm, respectively. TDCSPPMnCl was rapidly oxidized by NaCIO at pH 10. Addition of NaCIO (lxl0" 7 mole) to TDCSPPMnCl (3x10"* mole in 3 mL 0.1 M phosphate buffer) gave an intermediate 17 which absorbs at 446 nm. The newly formed 446 nm peak decreased slowly in intensity and at the same time shifted slowly to 432 nm as shown in Figure 2.7. The 472 nm peak also appeared during the process and increased in intensity. The 472 nm peak of TDCSPPMnCl was restored completely in about 19 hours. The UV-vis spectra of TDCSPPMnCl, 16, and 17 are shown in Figure 2.8. In the presence of veratryl alcohol, the oxidized intermediates of TDCSPPMnCl were observed less readily than in its absence. Intermediate 16 was not observed at pH 2 and pH 3 in the presence of veratryl alcohol (1x10 s mole). The oxidation of veratryl alcohol was observed under these conditions. The oxidized intermediate 16 and/or 17, therefore, was indeed formed but rapidly reduced by veratryl alcohol. At pH 7 and pH 10, the intermediate 16 was observed by UV-vis spectroscopy but 17 was not seen, suggesting that 17 was more reactive than 16. Together with the observation that 17 decomposed to give 16 and TDCSPPMnCl, it is suggested here that 17 is probably the oxomanganese(V) complex and 16 is probably the oxomanganese(IV) complex. It has been reported14 that the oxomanganese(V) porphyrin was much more reactive than the oxomanganese(TV) porphyrin for alkene epoxidation. The redox cycle of TDCSPPMnCl is shown in Scheme 2.1. As shown in Figure 2.8, T D C S P P M n ^ O (16) and TDCSPPMn v =0 (17) have distinctively different UV-vis spectra. It has been reported2,15 that the oxomanganese(rV) and oxomanganese(V) complexes of meso-tetramesitylporphyrin or meso-tetra(4-sulfonatophenyl)porphyrin have similar UV-vis spectra. The electron-withdrawing chloro substituents of TDCSPPMnCl might be responsible for this 57 TDCSPPMnv=0 (17) T D C S P P M n m C l ^ = > TDCSPPMn^O (13) STANDING OR VA (16) Scheme 2.1 The redox cycle of TDCSPPMnCl (VA: veratryl alcohol) difference. B. Oxidation of T D C S P P M n C l by H 2 0 2 , t -BuOOH, and mCPBA Hydrogen peroxide and t-BuOOH seem to be unable to oxidize TDCSPPMnCl below pH 7 under the conditions described above for NaCIO, since neither spectral change of the porphyrin nor increase in absorption at 310 nm (the oxidation of veratryl alcohol) was observed. At pH 10, both oxomanganese(V) and oxomanganese(IV) complexes were observed by UV-vis spectroscopy. mCPBA oxidized TDCSPPMnCl between pH 2 to pH 7 in a way similar to NaCIO. At pH 10, a mixture of TDCSPPMn v =0 (17) and T D C S P P M n 1 ^ (16) was observed. 58 300 400 500 BOO 700 WAVELENGTH Figure 2.7 The decay of the oxidized intermediate 17 (TDCSPPMn v=0, X ln„ 446 nm) to give intermediate 16 (Xma 432 nm) and the manganese(III) porphyrin ( X ^ 472 nm) in pH 10 buffer at room temperature 1.400 -i Ul u z < m cc o tn m < 1.133-0.865-0.598-0.331 -TDCSPPMif^Cl 0.063-r TDCSPPMnv=0 TDCSPPMn^O 300 400 500 600 700 WAVELENGTH Figure 2.8 The UV-vis spectra of TDCSPPMnCl (A^„ 472 nm, in pH 7 buffer), T D C S P P M n 1 ^ (16, Ani„ 432 nm, in pH 7 buffer), and TDCSPPMn v=0 (17, A to„ 446 nm, in pH 10 buffer) 2.1.2.2 Oxidized intermediates of Cl16TSPPMnCl A. Oxidation of Cl16TSPPMnCI by NaCIO ClieTSPPMn^O (19) *Cl1 6TSPPMnn (18) Scheme 2.2 The redox cycle of Cl 1 6 TSPPMnCl (VA: veratryl alcohol) Addition of NaCIO (lxlO" 7 mole) to Cl 1 6 TSPPMnCl (3X10"8 mole) in pH 7 buffer (3 mL, 0.1 M phosphate) resulted in the decrease of the 482 nm peak of Cl 1 6 TSPPMnCl and formation of two new peaks at 428 nm and 452 nm. Both new peaks decreased rapidly in intensity and the 482 nm peak was restored. Oxidation of Cl 1 6 TSPPMnCl (3x10"* mole) by more NaCIO (1x10^ mole) at pH 7 also gave peaks at 428 nm and at 452 nm. Addition of lxlO" 5 mole more of NaCIO resulted in an 61 intensity increase of the 428 nm peak and decrease of the 452 nm peak. When veratryl alcohol (lxlO-5 mole) was added to the solution, the 428 nm peak decreased in intensity and the 452 nm peak increased at first, then the 452 nm peak decreased and 482 nm peak appeared. The 310 nm peak of veratraldehyde increased rapidly until the 482 nm peak appeared. Interestingly, when the 310 nm peak stopped increasing and the 482 nm peak nearly returned to its original intensity, the 482 nm peak started to decrease and finally gave a new peak at 452 nm as shown in Figure 2.9. This mysterious intermediate (18) formed after veratryl alcohol oxidation had stopped (310 nm peak stopped increasing) was fairly stable and decayed to give the 482 nm peak of Cl16TSPPMnCl after overnight. It was inactive towards veratryl alcohol. Oxidation of Cl16TSPPMnCl (3X10"8 mole) by NaCIO (3x10s mole) in pH 7 buffer (3 mL) gave a intermediate 19 which has its Soret band at 428 nm. The 428 nm peak decreased in intensity slowly at room temperature in air. Intermediate 19 oxidized veratryl alcohol rapidly, the 428 nm peak decreased in intensity and a peak at 452 nm appeared (Figure 2.10). After that, the 482 nm peak also appeared and its intensity increased rapidly, the 452 nm peak and the 428 nm peak disappeared (Figure 2.11). The inactive intermediate 18 appeared again after the 482 nm peak was restored to its original intensity. Oxidation of Cl16TSPPMnCl (3X10"8 mole) by NaCIO (1x10* mole) at pH 10 (3 mL phosphate buffer) gave a species 20 which has its Soret peak at 452 nm. This species was stable and decayed to give the 482 nm peak of Cl16TSPPMnCl overnight. When veratryl alcohol was added to a solution containing 20, the 452 nm peak decreased rapidly to give a peak at 482 nm (Figure 2.12). After the 310 nm peak of veratraldehyde stopped increasing and the 482 nm peak increased to its original intensity, again, the inactive intermediate 18 was formed. Though having their Soret 62 0.0000 300 4 0 0 500 600 700 WAVELENGTH Figure 2.9 The conversion of Cl 1 6 TSPPMnCl (Xm„ 482 nm) to the intermediate 1 8 (Cl 1 6 TSPPMn n , 452 nm) in pH 7 phosphate buffer at room temperature after veratryl alcohol oxidation stopped figure 2.10 500 600 v e r a t r y j alcohol 700 Figure 2.11 The reduction of intermediate 20 (X^„ 452 nm) to Cl 1 6 TSPPMnCl 482 nm) by veratryl alcohol in pH 7 buffer at room temperature Figure 2.12 The reduction of intermediate 20 (A*,„ 452 nm) to Cl 1 6 TSPPMnCl ( A ^ 482 nm) by veratryl alcohol in pH 10 buffer at room temperature bands at the same position, the inactive intermediate 18 and the one (20) which can oxidize veratryl alcohol have different extinction coefficients as shown in Figure 2.13. No spectral changes were observed when NaCIO was added to Cl 1 6 TSPPMnCl at pH 2 or pH 3. It is proposed here that intermediate 19 which is very reactive and decays to give the intermediate 20 is Cl 1 6 TSPPMn v =0. The intermediate 20 which is able to oxidize veratryl alcohol is probably CljjTSPPMn™^. The UV-vis spectrum of manganese(UI), oxomanganese(IV) (20), and oxomanganese(V) (19) porphyrins are shown in Figure 2.14. The inactive intermediate 18 was found to be Cl 1 6TSPPMn(II) as rationalized below. As this intermediate was unable to oxidize veratryl alcohol but was observed only in the presence of veratryl alcohol, it must be a product of the reaction between the manganese(III) porphyrin and veratryl alcohol or one of the oxidation products of veratryl alcohol. It was noted that at pH 10, the UV-vis spectrum of Cl 1 6 TSPPMnCl had a shoulder around 452 nm. Addition of sodium hydroxide to the solution shifted the 482 nm peak of Cli 6 TSPPMnCl to 456 nm. This 456 nm species was believed to be a hydroxide adduct of the manganese(III) porphyrin.2 It had a similar UV-vis spectrum to that of the inactive intermediate 18. Addition of hydrochloric acid to the 456 nm species shifted the Soret band back to 482 nm instantly. The inactive intermediate 18, however, did not change back to 482 nm upon the addition of hydrochloric acid. Addition of a large excess of veratryl alcohol (3x10 s mole) to the manganese(III) porphyrin (3x10"* mole) at pH 10 resulted in a very slow decrease of the 482 nm peak of Cli 6TSPPMnCl and a correspondingly slow formation of a new peak at around 450 nm. The shifting from 482 nm to 450 nm was not complete even 67 0 .0000 -\ . . . . 1 . . • . 1 . . . • 1 i • • i 300 400 500 600 700 WAVELENGTH Figure 2.13 The UV-vis spectra of intermediate 18 (Cl I 6 TSPPMn n , in p H 10 buffer) and 20 (Cl 1 6 TSPPMrr=0, in p H 10 buffer) Figure 2.14 The UV-vis spectra of Cl 1 6 TSPPMnCl 482 nm, in pH 7 buffer), Cl 1 6 TSPPMn v =0 (19, 428nm, in pH 7 buffer), and Cl 1 6TSPPMn I V=0 (20, A*. 452 nm, in pH 10 buffer) after standing overnight The formation of the 450 nm species, which was probably a veratryl alcohol-porphyrin complex, was much slower than the formation of the inactive intermediate 18. Addition of veratraldehyde (3xl0"5 mole), the major product of veratryl alcohol oxidation, to Cl16TSPPMnCl (3x10* mole) at pH 10 (3 mL 0.1 M phosphate buffer) did not change the UV-vis spectrum of the manganese(HI) porphyrin. It has been reported that oxidation of veratryl alcohol by lignin peroxidase produced superoxide.18 Manganese porphyrins could also catalyze the oxidation of veratryl alcohol and give superoxide. Reaction of superoxide with the manganese(in) porphyrin can give a manganese(IH)-superoxide complex, which is unstable at room temperature and decomposes to give dioxygen and the manganese(U) porphyrin as shown below: Cl16TSPPMn(ffl)Cl + 02" -> Cl16TSPPMn(m)-02- -> Cl16TSPPMn(n) + 0 2 Reactions of manganese(m) porphyrins and superoxide in organic solvents to give manganese(U) porphyrins have been reported.15,19 Manganese porphyrins were found to bind dioxygen reversibly at -78°C. At room temperature, however, the manganese(II) porphyrin was rapidly oxidized by dioxygen.20 The fact that Cl16TSPPMn(U) was relatively stable at room temperature suggests that it might be a much better haemoglobin model for reversible dioxygen binding than other simple manganese porphyrins previously studied. Cl16TSPPMn(U) (18) was oxidized rapidly to give the manganese(III) porphyrin by adding excess NaCIO, explaining the fact that Cl16TSPPMn(II) (18) was observed 70 only after the oxidation of veratryl alcohol had stopped, which happened when all the NaCIO was consumed. Before the NaCIO was depleted, Cl 1 6TSPPMn(U) could also be formed by the reaction described above. It could not be observed, however, by UV-vis spectroscopy due to its rapid oxidation by NaCIO. In order to confirm that the inactive intermediate 18 was Cl 1 6TSPPMn(II), the manganese(U) porphyrin was made by another method to compare its UV-vis spectrum with that of 18. Addition of small amounts of solid sodium dithionite to Cl 1 6 TSPPMnCl (3X10"8 mole in 3 mL 0.1 M pH 7 phosphate buffer) gave a species which had its Soret band at 450 nm. This species, however, was not stable and decomposed rapidly to give an orange solution. This experiment was also very difficult to repeat since excess sodium dithionite destroyed the porphyrin instandy to give an orange solution. Experiments carried out in pH 10 phosphate buffer (0.1 M) gave similar results. Reduction of Cl 1 6 TSPPMnCl (3x10"* mole) by excess sodium borohydride in pH 10 phosphate buffer (3 mL) gave a complex which had its Soret band at 450 nm and another small peak at 582 nm. The UV-vis spectrum of this complex was the same as that of 18 except a 2 nm wavelength difference between the two Soret bands. Because the different solvent condition, these two spectra can be considered as the same. The formation of Cl 1 6TSPPMn(U) by reacting the manganese(III) porphyrin and superoxide was also demonstrated. Aliquots of potassium superoxide dissolved in acetonitrile containing 18-crown-6 ether were added to a Cl 1 6 TSPPMnCl solution (3xl0 8 mole in 3 mL pH 10 phosphate buffer) until the 452 nm peak reached its maximal absorption. Further addition of potassium superoxide decreased the intensity of the 452 nm peak. It was found that the 452 nm peak thus formed had a smaller intensity compared with that of the manganeseQT) porphyrin formed from the reaction of 71 Cl1 6TSPPMnCl and NaCIO in the presence of veratryl alcohol when the initial concentrations of the manganese(III) porphyrin were the same. It is possible that hydrogen peroxide, formed from potassium peroxide (which was present in potassium superoxide), might have interfered with the reaction. Addition of veratryl alcohol could reduce the oxidized intermediates of the manganese porphyrin, which were produced by hydrogen peroxide oxidation, to regenerate the manganese(TJj) porphyrin. Reaction of manganese(m) rxMphyrin with superoxide may produce pure manganese(JJ) porphyrin. Indeed, reaction of Cl1 6TSPPMnCl (3x10s mole in 3 mL pH 10 buffer) with potassium superoxide in the presence of veratryl alcohol (1x10s mole) gave a 452 nm peak which had similar intensity as that of the manganese(II) porphyrin formed after the oxidation of Cl1 6TSPPMnCl by NaCIO in the presence of veratryl alcohol. While no strong evidence was available to support the formation of superoxide during the oxidation of veratryl alcohol by Cl1 6TSPPMnCl, the effect of a superoxide dismutating agent on the formation of Cl16TSPPMn(II) would provide indirect evidence that superoxide was involved. Manganese(U) complexes are well known model systems of superoxide dismutase.21,22 The effect of manganese(II)-lactate on the formation of Cl16TSPPMn(U) was studied. In the presence of veratryl alcohol (1x10s mole) and manganese(II)-lactate (ImM and 60 mM concentration, respectively, in 3 mL pH 7 buffer), reaction of Cl1 6TSPPMnCl (3x10* mole) and NaCIO (2x10s mole) did not give the manganese(U) porphyrin, suggesting that the superoxide produced was effectively dismutated by manganese(II)-lactate. In the absence of manganese(IJ)-lactate and under otherwise the same condition, the manganese(U) porphyrin was observed by UV-vis spectroscopy. 72 B. Oxidation of Cl 1 6TSPPMnCl by mCPBA Unlike NaCIO, mCPBA was able to oxidize Cl 1 6 TSPPMnCl over a wide range of pH. The oxidized intermediates at pH 2 and pH 3 were found to have their Soret bands at slighdy different wavelengths compared with those formed by using NaCIO as oxidant. When mCPBA (5x10-* mole) was added to Cl 1 6 TSPPMnCl (3x10"* mole) in 3 mL 0.1 M pH 2 phosphate buffer, the UV-vis spectrum showed a broad peak at 420 nm which had a shoulder at 474 nm (Figure 2.15). Further addition of mCPBA decreased the intensity of both peaks. Addition of veratryl alcohol ( lxlO' 5 mole) brought the Soret peak back to 482 nm rapidly. A peak at 420 nm and another peak at 460 nm were found in the UV-vis spectrum when mCPBA (1X10"* mole) was added to Cl 1 6 TSPPMnCl (3x10"* mole) at pH 3 (3 mL 0.1 M phosphate buffer). The 460 nm peak decreased in intensity and the 420 nm peak remained almost unchanged upon further addition of 5x10* mole of mCPBA. The intensity of both peaks decreased when the solution was allowed to stand at room temperature. When the reactions were carried out in phosphate buffer of pH 6^  or higher, the oxidation of Cl 1 6 TSPPMnCl by mCPBA was similar to that using NaCIO as oxidant. Oxomanganese(V) and oxomanganese(IV) porphyrin have their Soret absorption at 430 nm and 454 nm, respectively. Cl 1 6TSPPMn(U) was formed at above pH 5 during the oxidation of veratryl alcohol by Cl 1 6 TSPPMnCl and mCPBA. The manganese(II) porphyrin was formed very slowly at pH 5. Superoxide formed from the oxidation of veratryl alcohol probably decomposed rapidly at pH 2 and pH 3 and therefore the manganese(II) porphyrin was not formed. 73 —1 0 . 0000 -\ . .——. . 1 . . * • 1 . • • • 1 • • • ' 1 300 400 500 600 700 W A V E L E N G T H Figure 2.15 The oxidation of Cl16TSPPMnCl (3x10" mole) by mCPBA (5X10"6 mole) in pH 2 phosphate buffer at room temperature ) 2.1.2.3 Oxidized intermediates of TDCSPPFeCl Oxidation of TDCSPPFeCl (3x10-* mole) with NaCIO (lxlO"7 mole) in pH 7 phosphate buffer (3 mL, 0.1 M) shifted the Soret band from 424 nm to 430 nm. Additional NaCIO (lxlO-6 mole) further shifted the Soret band to 431 nm. The 431 nm peak shifted back to 424 nm slowly upon standing at room temperature. When NaCIO (lxlO"6 mole) was added to TDCSPPFeCl (3x10-* mole in 0.1 M phosphate buffer) at pH 10, the UV-vis spectrum showed a peak at 432 nm (Figure 2.16). Further addition of NaCIO caused no change in the spectrum. The 432 nm peak was also observed when TDCSPPFeCl (3x10"* mole) was oxidized by NaCIO (lxlO"6 mole) at pH 12 (3 mL, 0.1 M borate buffer). The formation of oxidized intermediates of TDCSPPFeCl was not observed at pH 2 and pH 3. The oxidation of TDCSPPFeCl by mCPBA gave similar spectral changes. The nature of the species (21) which has its Soret band at 432 nm is not clear. 2.1.2.4 Oxidized intermediates of Cl16TSPPFeCl The Soret band of Cl16TSPPFeCl (3x10"* mole) decreased in intensity when NaCIO (lxlO-6 mole) was added to the iron porphyrin at pH 2 or pH 3 (3 mL phosphate buffer). When NaCIO (1x10"* mole) was added to Cl16TSPPFeCl (3x10* mole) at pH 7 or pH 10 (3 mL 0.1 M phosphate buffer), the Soret band of the iron (III) porphyrin decreased in intensity and a new peak at 448 nm was formed. The 422 nm peak did not disappear completely even in the presence of a large excess (3xl0'5 mole) of NaCIO. When Cl16TSPPFeCl (3x10"* mole in 3 mL 0.1 M phosphate buffer) was oxidized by NaCIO (1x10s mole) at pH 12 (3 mL 0.1 M borate buffer), a peak at 448 75 nm with a shoulder at 422 nm was formed. The UV-vis spectra of Cl 1 6 TSPPFeCl and that of the oxidized intermediate (22) are shown in figure 2.17. A dioxygen-iron(H) porphyrin complex, Cl 1 6TSPPFe(II)-0 2 (23) was observed when Cl 1 6TSPPFeCl was oxidized by NaCIO under basic condition (pH£7) at room temperature in the presence of veratryl alcohol. When NaCIO (5x10"* mole) was added to Cl 1 6 TSPPFeCl (3x10* mole in 3 mL 0.1 M pH 7 phosphate buffer) in the presence of veratryl alcohol (1x10 s mole), the 422 nm peak decreased in intensity and the 448 nm peak appeared and increased in intensity. The 310 nm peak of veratraldehyde kept increasing until the 448 nm peak disappeared and the 422 nm peak was restored. The 422 nm peak decreased in intensity after that and gave the dioxygen-iron(II) porphyrin complex which had its Soret band at 446 nm (Figure 2.18). The presence of manganese(II)-lactate, a superoxide dismutating agent, inhibited the formation of this dioxygen-iron(II) porphyrin complex, suggesting that the dioxygen complex was formed from the reaction of superoxide and the iron(UI) porphyrin: Cl1 6TSPPFeCl + 0 2 -> Cl16TSPPFe(II)-02 The dioxygen complex (23) reacted with excess NaCIO to give a UV-vis spectrum characteristic of the iron(ITf) porphyrin, explaining the fact that the dioxygen complex was observed only after the oxidant (NaCIO) was consumed. The dioxygen-iron(II) porphyrin complex (23) reacted with carbon monoxide to give the CO-iron(II) porphyrin complex (Figure 2.19), which showed the same UV-vis spectrum as that of the complex prepared by reacting Cl 1 6 TSPPFeCl with excess sodium dithionite at pH 7 followed by treating the solution with carbon monoxide. 77 0 .0000 -\ . . 1 . , . .——. . r . . . . , . . . . 1 300 400 500 600 700 WAVELENGTH Figure 2.17 The oxidation of Cl)6TSPPFeCl (k^ 422 nm, 3X10"8 mole) to the A*. 448 nm intermediate (22) by NaCIO (1x105 mole) in pH 12 borate buffer (3 mL) at room temperature Figure 2.18 The UV-vis spectra of Cl 1 6TSPPFeCl (A*. 422 nm) and Cl 1 6 TSPPFe n -0 2 (23, 446 nm) in pH 7 phosphate buffer (0.1 M) Figure 2.19 The reaction of CO and Cl 1 6TSPPFe"-0 2 (23) formed after the oxidation of veratryl alcohol to give Cl 1 6TSPPFe n-CO 440 nm) in pH 7 phosphate buffer OigTSPPFeCl (3x10"* mole) also reacted with potassium superoxide in the presence of veratryl alcohol (lxlO' 5 mole) to give the dioxygen-irontTT) porphyrin complex 23. Cl16TSPPFeCl destruction by hydrogen peroxide was observed even in the presence of a large excess of veratryl alcohol. No spectral change was observed when Cl16TSPPFeCl (3x10-" mole) was treated with t-BuOOH (5x10* mole) at pH 2 and pH 3. The reaction of Cl16TSPPFeCl with t-BuOOH at pH 7 or pH 10 was similar to that with NaCIO. No oxidized intermediates were observed when Cli6TSPPFeCl was oxidized by mCPBA at pH 2 and pH 3, pH 5 was the minimum pH to observe the oxidized intermediate 22 (X^ 448 nm) and the dioxygen-iron(II) porphyrin in the presence of veratryl alcohol. 2.1.3 Factors affecting the catalytic activity of the metalloporphyrins Oxidation of veratryl alcohol by Ugnin peroxidase gave veratraldehyde as a major product together with small amounts of quinones and aromatic ring cleaved products. The oxidation of veratryl alcohol by protohemin, TPPFeCl, or TDCSPPFeCl gave similar products as wiU be described in Chapter 3. Veratryl alcohol was used as a substrate in this experiment and the formation of veratraldehyde was used as a criterion for comparing the catalytic activities of metaUoporphyrins under different conditions. Veratraldehyde has a strong absorption at 310 nm and its formation can be conveniendy foUowed by using a rapid scanning Diode Array UV-vis spectrometer. 81 2.1.3.1 pH of the solvent A. TDCSPPFeCl as catalyst The pH dependency of the oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA is shown in Figure 2.20. Both the rate and yield of veratraldehyde formation were highly dependent on the pH of the buffer solution. It can be seen from Figure 2.20 that at pH 3 and pH 5, the reaction stopped soon after addition of mCPBA and the final yield of veratraldehyde was lower than that at pH 2. It was found that the reaction stopped because the oxidant was consumed. The porphyrin catalyst was still active after the reaction and addition of another aliquot of mCPBA restarted the reaction as shown in Figure 2.21. The oxidation of veratryl alcohol (1x10* mole) by TDCSPPFeCl (3x10-* mole) and NaCIO (5x10* mole) in 0.1 M aqueous buffer of various pH's is shown in Figure 2.22. The highest rate was seen at pH 5 and the highest yield was achieved at pH 7. Little reaction was observed at pH 2 and pH 3. The catalyst was also reactive after the reaction as shown in Figure 2.23. TDCSPPFeCl (3x10-* mole) catalyzed oxidation of veratryl alcohol (1x10* mole) in 0.1 M aqueous buffer of various pH's using t-BuOOH as oxidant is shown in Figure 2.24. There were no observable reactions at pH 8 and pH 10. There seemed to be an induction period at pH 2, 3, 5, and 7. It has been suggested that the displacement of chloride by mCPBA was a slow process and was responsible for the induction period in the oxidation of mgso-tetramesitylporphyrin iron chloride by mCPBA.6 An induction period was observed in this laboratory23 for the oxidation of we^ o-tetra(2,6-dichlorophenyl)porphyrin iron chloride (TDCPPFeCl) but not for the corresponding 82 A • pH 2.0 2.50 " 2.00 " 0.0 1.0 2.0 3.0 Time (min.) Figure 2.20 The oxidation of veratryl alcohol (5x10* mole) by TDCSPPFeCl (1x10"* mole) and mCPBA (2x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5 and 6 buffers are 0.1 M citrate and all others are 0.1 M phosphate. 83 A E 1 .50-c Time (min.) F i g u r e 2.21 T h e o x i d a t i o n o f v e r a t r y l a l c o h o l (5x10* m o l e ) b y T D C S P P F e C l (1x10* m o l e ) a n d m C P B A i n 4 m L 0.1 M p H 5 c i t r a t e b u f f e r a t r o o m t e m p e r a t u r e , t h e s a m e a m o u n t o f m C P B A (2x10* m o l e ) w a s a d d e d a t t i m e = 0 a n d t i m e = 1 m i n . 0 . 5 0 0 r Time (seconds) Figure 2.22 The oxidation of veratryl alcohol ( 1 x 1 0 s mole) by T D C S P P F e C l ( 3x10* mole) and NaCIO (5x10^ mole) at room temperature in 4 m L aqueous buffer of various p H values, p H 5, and 6 buffers are 0.1 M citrate and all others are 0.1 M phosphate. 00 O N E c o 0) o c D »_ O OT j Q < 0 . 5 0 0 n 0 . 4 0 0 -0 . 3 0 0 -Time (seconds) Figure 2.23 The oxidation of veratryl alcohol (1x10 s mole) by TDCSPPFeCl (3x10* mole) and NaCIO in 4 mL 0.1 M pH 6 citrate buffer at room temperature, NaCIO (5x10* mole) added at time = 0 and time = 90 seconds 0.500 pH 6 oo E c o T CD U c o JD o cn X) < 0.400-0.300-0.200-0.100 0.000 ,A .9' 0 60 120 180 240 Time (seconds) 300 ,A P H 5 pH 2 & pH 7 360 Figure 2.24 The oxidation of veratryl alcohol (1x10 s mole) by t-BuOOH (5x10* mole) and TDCSPPFeCl (3xl(r* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 2, 3, and 7 buffers are 0.1 M phosphate, pH 5 and 6 buffers are 0.1 M citrate. hydroxide, TDCPPFeOH. A/e5c9-tetra(2,6-dichloro-3-sulfonatophenyl)porphyrin iron hydroxide (TDCSPPFeOH) was synthesized and tested for the presence of induction periods during the oxidation of veratryl alcohol. As shown in Figure 2.25 (curve 1), the induction period still exists in the oxidation of veratryl alcohol by TDCSPPFeOH and t-BuOOH at pH 3. Addition of sodium chloride, which will shift the equilibrium: TDCSPPFeOH + Cl ^ = TDCSPPFeCl + OH to the right, not only failed to extend the induction period but speeded up the reaction as shown in Figure 2.25. This result indicates that chloride as axial ligand was not responsible for the induction period. The stimulating effect for the oxidation of veratryl alcohol caused by sodium chloride is probably a simple ion effect. Lignin peroxidase and biomimetic catalysts oxidize lignin model compounds through a cation radical mechanism. Increasing ionic strength will stabilize the cation radical intermediates. The rate of the oxidation of veratryl alcohol increased with increasing concentration of sodium chloride (Figure 2.25). As shown in Figure 2.26, addition of 1 M sodium sulfate results in a 4 fold increase of the reaction rate. Not all inorganic salts increase the reaction rate. The presence of 1 M sodium fluoride or sodium nitrate (in 3 mL 0.1 M pH 3 phosphate buffer) completely inhibited the oxidation of veratryl alcohol ( lx lO' 5 mole) by t-BuOOH (lxlO- 5 more) and TDCSPPFeOH (3X10"8 mole). The effect of buffer concentration on the oxidation of veratryl alcohol was also studied. As shown in Figure 2.27, increasing buffer concentration also increases the rate of the oxidation of veratryl alcohol by TDCSPPFeOH and t-BuOOH. 88 0.500 oo E c o <U o c o .O o co < 0.400 -0.300 0.200 0.100 0.000 o 60 JL -o- -o-=6 . 4 3 5 1 120 180 240 300 Time (seconds) 360 Figure 2.25 The oxidation of veratryl alcohol ( lx lO' 5 mole) by TDCSPPFeOH (3x10* mole) and t-BuOOH (1x10 s mole) at room temperature in 4 mL 0.1 M pH 3 phosphate buffer containing various concentrations of sodium chloride (M), 1, 0; 2, 0.1; 3, 0.2; 4, 0.5; 5, 1.0; 6, 2.0 o E c o <u o c D . Q i _ o X) < 0.700 r 0.600 0.500 0.400 0.300 0.200 0.100 0.000 - o — ° _o—°" 0 60 120 180 240 300 Time (seconds) 360 Figure 2.26 The oxidation of veratryl alcohol (1x10 5 mole) by TDCSPPFeOH (3x10"* mole) and t-BuOOH ( l x l O 5 mole) at room temperature in 4 mL 0.1 M pH 3 phosphate buffer containing: 1, 1 M Na 2S0 4; 2, 1 M NaCl; 3, no other inorganic salts VO E c o <D O c D o (n JD < 0.400 0.350 0.300 0.250 0.200 0.150 0.100 0.050 0.000 A n A' . - _ -O' X T -o-X T ja 2 0 60 120 180 240 300 Time (seconds) 360 Figure 2.27 The oxidation of veratryl alcohol (1x10 s mole) by TDCSPPFeOH (3x10"* mole) and t-BuOOH (1x10 s mole) at room temperature in 4 mL pH 3 phosphate buffer of various concentrations (M), 1, 0.1; 2, 0.5; 3, 1.0 Although purified by vacuum distillation, veratryl alcohol was shown by GC analysis to contain a trace amount of methyl 4-hydroxy-3-methoxybenzoate, which was responsible for the induction period. It was reported that the same compound delayed the oxidation of veratryl alcohol by lignin peroxidase.24 Addition of methyl 4-hydroxy-3-methoxybenzoate (lxlO"6 mole) dramatically slowed down the oxidation of veratryl alcohol (1x10s mole in 3 mL 0.1 M pH 3 phosphate buffer) by TDCSPPFeOH (3x10* mole) and t-BuOOH (lxlO"5 mole). The induction period was not seen when veratryl alcohol was purified by double distillation. Hydrogen peroxide seemed to be an inefficient oxidant for TDCSPPFeCl catalyzed veratryl alcohol oxidation. The reaction was slow at every pH (Figure 2.28). B. CI 1 6 TSPPFeCl as catalyst Figure 2.29 shows the pH dependency of Cl16TSPPFeCl catalyzed oxidation of veratryl alcohol by mCPBA. The maximal catalytic activity was observed at pH 2 and the reaction was slower and the yield was lower at higher pH. The catalyst was still active after the reaction (Figure 2.30). Cl16TSPPFeCl (3x10* mole) and t-BuOOH (5x10* mole) oxidized veratryl alcohol (lxlO"5 mole) effectively only in pH 5, 6, and 7 buffers (3 mL) as shown in Figure 2.31. The reaction was very slow at pH 2, 3, 8, and 10. Cl I 6TSPPFeCl catalyzed oxidation of veratryl alcohol was favoured at high pH when NaCIO was the oxidant (Figure 2.32). Cl16TSPPFeCl was not stable towards hydrogen peroxide and the oxidation of veratryl alcohol (lxlO5 mole) by Cl16TSPPFeCl (3x10* mole) and H A (5x10* mole) was very slow. 92 VO E. c o cu o c o X) o co JD < 0.200 0.150 0.100 0.050 y° PH 2 o D O / V o ^ n - D ^ ^ D — D — D — D p H 3 p H 10 p H 5 PH 6 0.000 0 30 60 90 120 150 180 Time ( s e c o n d s ) Figure 2.28 The oxidation of veratryl alcohol (1x10 s mole) by TDCSPPFeCl (SxlO"8 mole) and H 2 0 2 (5x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 2, 3 and 10 buffers are 0.1 M phosphate, pH 5 and 6 buffers are 0.1 M citrate. T i m e ( m i n . ) Figure 2.29 The oxidation of veratryl alcohol (5x10* mole) by Cl 1 6 TSPPFeCl (1x10* mole) and mCPBA (2x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5, and 6 buffers are 0.1 M citrate and all others are 0.1 M phosphate. 94 E c o ro <D o c o X) l_ o 0) XI < 1.200 1.000 0.800 0.600 0.400 0.200 A 0.000 4 / / / -o—O-—o / / / / / 0 30 60 90 120 150 180 Time (seconc Figure 2.30 The oxidation of veratryl alcohol ( lxl f r 5 mole) by Cl I 6 TSPPFeCl (1x10* mole) and mCPBA in 4 mL 0.1 M pH 5 citrate buffer at room temperature, mCPBA (lxlO" 6 mole) added at time = 0 and time =90 seconds E c o cu u c D JD i _ O CO JD < 0.700-j 0.600-0.500-0.400 0.300-1 0.200 0.100 0.000 0 y. / yo PH5 / ° v PH 6 y y y sy _ A—A-r~ -A—A-pH 7 60 120 180 240 Time (seconds) 300 360 Figure 2.31 The oxidation of veratryl alcohol (1x10 s mole) by Cl 1 6 TSPPFeCl (3x10* mole) and t-BuOOH (5x10* mole) in 4 mL 0.1 M phosphate buffer (pH 7) or 0.1 M citrate buffer (pH 5 and 6) at room temperature A 0.80" 0 30 60 90 120 150 180 T i m e ( s e c o n d s ) Figure 2.32 The oxidation of veratryl alcohol ( l x l C 5 mole) by Cl 1 6 TSPPFeCl (3x10* mole) and NaCIO (5x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 7 and 10 buffers are 0.1 M phosphate, pH 5 and 6 buffers are 0.1 M citrate, and pH 9.2 buffer is 0.1 M borate. 97 C. TDCSPPMnCl as catalyst TDCSPPMnCl catalyzed oxidation of veratryl alcohol by various oxidants was also highly dependent on the pH of the solvent as shown in Figures 2.33, 2.34, and 2.35. There was no observable loss of catalytic activity of the manganese porphyrin during the reaction. As shown in Figure 2.36, when the oxidant was consumed and the 310 nm peak stopped increasing, addition of a second aliquot of oxidant restarted the reaction and caused the same increase of 310 nm absorption. When t-BuOOH (5x10"* mole) was the oxidant, TDCSPPMnCl (3x10"* mole) catalyzed oxidation of veratryl alcohol (1x10 s mole) was very slow at pH 10 and no reaction was observed between pH 3 and pH 8. D. Cl16TSPPMnCl as catalyst Cl 1 6 TSPPMnCl catalyzed oxidation of veratryl alcohol was favoured at high pH when NaCIO or H 2 0 2 was the oxidant as shown in Figures 2.37 and 2.38. When mCPBA was the oxidant, the catalytic activity order was pH 3 > pH 2 > pH 5 > pH 8, pH 10, pH 7 (Figure 2.39). When t-BuOOH (5x10* mole) was the oxidant, the reaction (3X10-8 mole Cl 1 6 TSPPMnCl, 1x10 s mole veratryl alcohol in 3 mL buffer) was very slow between pH 5 and pH 10 and no reaction was observed at pH 2 and pH 3. As discussed earlier, Cl1 6TSPPMn(JJ) was produced at high pH from the reaction of the manganese(III) porphyrin and superoxide. Oxidant could be consumed in vain for oxidizing the manganese(II) porphyrin back to the manganese(III) porphyrin. It is known that the dismutation of superoxide in aqueous solution is inversely proportional to the pH. More oxidant would be consumed at higher pH by oxidizing the 98 VO VO E c o U c D JD i_ O (fi JD < 2.000 1.500-1.000 0.500 0.000 ^1=1=•=! ,A -A' -A A — A — A — A p H 3 p H 5 p H 2 p H 6 p H 7 p H 8, 10 0 30 60 90 120 Time (seconds) 150 180 Figure 2.33 The oxidation of veratryl alcohol (5x10* mole) by TDCSPPMnCl ( l xHr 8 mole) and mCPBA (2x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5 and 6 buffers are 0.1 M citrate and all others buffers are 0.1 M phosphate. E c o ro < u o c o JD o t o JD < 1.000 pH 9.2 150 180 Time (seconds) Figure 2.34 The oxidation of veratryl alcohol (1x10 s mole) by TDCSPPMnCl (3x10* mole) and H 2 0 2 (5x10* mole) in 4 mL 0.1 M pH 10 buffer (phosphate) or pH 9.2 buffer (borate) at room temperature E c o CD o c o - Q o J Q < 0.500 i 0.400 0.300-0.200 0.100 0.000 -XJ" _ • — • — o — • — • — • pH 6 • • • /P K A A A A A A pH 7 pH 5 pH 8 A _ A — A — A — A — A — A -• V A — A — A pH 2 S ± 8 ^ 8 ^ 8 = 8 = 8 = 8 = 8 = 8 = 8 = 8 = 8 = 8 = = e = 8 = 8 = 8 p H 1 0 ' — - v — v — y — \ 7 — 1 7 — y — v — v — y — v — v — y — V — g = y _ pH 3 0 30 60 90 120 150 180 T ime ( s e c o n d s ) Figure 2.35 The oxidation of veratryl alcohol (1x10 s mole) by TDCSPPMnCl (3x10* mole) and NaCIO (5x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5 and 6 buffers are 0.1 M citrate and all others are 0.1 M phosphate. o to E c o ro d) o c o X) o CO J Q < 2.200-2.000 1.800-1 1.600 1 . 4 0 C H 1.200 1.000 0.800 0.600 0.400 0.200 v-o—o / / / / 0.000 * 0 60 120 180 240 300 360 420 480 Time (seconds) Figure 2.36 The oxidation of veratryl alcohol (5x10* mole) by TDCSPPMnCl (1x10" mole) and mCPBA in 4 mL 0.1 M pH 5 citrate buffer at room temperature, mCPBA (2x10* mole) added at time = 0 and time = 240 seconds 1.20 a 2' 0.80-C3 O a as x> u o n < 0, 0.40 A-pH 7.0 n- D pH 8.0 _A—A—A-A-^-A—A-A—rj^fi—A^-A pH 5.0 ^ P H 9 ' 2 100 150 200 Time (seconds) Figure 2.37 The oxidation of veratryl alcohol ( l x l ( j 5 mole) by Cl 1 6 TSPPMnCl (3x10"* mole) and NaCIO (5x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5 and 6 buffers are 0.1 M citrate, pH 7, 8, and 10 buffers are 0.1 M phosphate, and pH 9.2 buffer is 0.1 M borate. 103 A Time (second) Figure 2.38 The oxidation of veratryl alcohol ( l x l O 5 mole) by Cl 1 6 TSPPMnCl (3x10"* mole) and H 2 0 2 (5x10* mole) in 4 mL 0.1 M pH 8 (phosphate), 9.2 (borate), and 10 (phosphate) buffer at room temperature A Time (min.) Figure 2.39 The oxidation of veratryl alcohol (5x10* mole) by Cl 1 6 TSPPMnCl (lxlO" 8 mole) and mCPBA (2x10* mole) at room temperature in 4 mL aqueous buffer of various pH values, pH 5 and 6 buffers are 0.1 M citrate and all other buffers are 0.1 M phosphate. 105 manganese(U) porphyrin. It has been shown that the oxidation of a metalloporphyrin by an oxidant was pH dependent. The rate of oxidation of iron meio-tetra(2,6-dimethyl-3-sulfonatophenyl)porphyrin (24) by hydrogen peroxide in aqueous buffer increases with increasing p H 2 5 (pH 1-12). The oxidation of manganese m«o-tetra(2,6-dimethyl-3-sulfonatophenyl)porphyrin (25) by hydrogen peroxide was also studied26 and the reaction rate increased with increasing pH in the pH range from 7.6 to 12.1. The rate of the oxidation of veratryl alcohol depends on two reactions: MPor + oxidant -» (MPor)oxidi2ed (1) (MPor)oxidized + veratryl alcohol -» MPor + veratraldehyde (2) where MPor represents metalloporphyrin and (MPor)O J d d i 2 e d is its oxidized intermediate. The rate of reaction (1) is a complex function of pH and it increases with increasing pH. 2 5 , 2 6 The fact that the oxidized intermediates of the iron and manganese porphyrins were observed by UV-vis spectroscopy at high pH but not at low pH in the presence of a large excess of veratryl alcohol, indicates that the rate of reaction (2) increases with decreasing pH. The optimum pH for the overall reaction will be determined by the kinetics of both reactions within the pH range of the study. The optimal pH at which the metalloporphyrins showed maximal catalytic activity for veratryl alcohol oxidation is summarized in Table 2.1. The oxidation of veratryl alcohol by Cl 1 6 TSPPMnCl/H 2 0 2 , Cl 1 6TSPPMnCl/t-BuOOH, TDCSPPMnCl/H 2 0 2 , or TDCSPPMnCl/t-BuOOH showed maximum rate at the highest pH used (pH 10), probably because reaction (1) was the rate limiting step in these systems. 106 Table 2.1 Optimum pH for the oxidation of veratryl alcohol by various metalloporphyrin/oxidant combinations at room temperature Metalloporphyrin Oxidant Optimal pH Cl 1 6 TSPPMnCl mCPBA 3 Cl 1 6 TSPPFeCl mCPBA 2 TDCSPPMnCl mCPBA 3 TDCSPPFeCl mCPBA 2 TDCSPPMnCl NaCIO 6 TDCSPPFeCl NaCIO 7 Cl 1 6 TSPPFeCl t-BuOOH 5 TDCSPPFeCl t-BuOOH 6 Cl 1 6 TSPPMnCl t-BuOOH 10 TDCSPPMnCl t-BuOOH 10 Cl 1 6 TSPPMnCl NaCIO 10 Cl 1 6 TSPPFeCl NaCIO 10 TDCSPPFeCl H 2 0 2 2 TDCSPPMnCl H 2 0 2 10 Cl 1 6 TSPPMnCl H 2 0 2 10 107 TDCSPPMnCl and Cl1 6TSPPMnCl have higher redox potentials than their coniesponding iron porphyrins,26 t-BuOOH and HJOJ are weaker oxygen donors than mCPBA. Both factors are responsible for a slow reaction (1). Iron and manganese porphyrins can mimic the functions of peroxidases as well as catalases at the same time. As a peroxidase model, the two-electron oxidized metalloporphyrin oxidizes substrates by one-electron oxidation. The two-electron oxidized metaUoporphyrin can also oxidize HJOJ to give 0 2 and FLO and thus function as a catalase. It was reported25 that the oxidation of hydrogen peroxide to oxygen by iron porphyrin 24 competed with its peroxidase like reaction and the former reaction was favoured with increasing pH. For the manganese porphyrin 25, just the opposite was true.26 The peroxidase activity of 25 was higher at higher pH and its catalase activity was higher at lower pH. It is, therefore, not surprising to find that TDCSPPFeCl showed its maximal activity under acidic condition (pH 2) for veratryl alcohol oxidation by H 20 2 . TDCSPPFeCl decomposed H^Oj more rapidly at higher pH and had litde peroxidase activity, veratryl alcohol was therefore oxidized slowly and inefficiendy at high pH. Better stability of the catalyst at low pH (Fig. 2.1, 2.2) was probably also a factor. All the reactions using mCPBA as oxidant are favoured at low pH. mCPBA is a good single oxygen donor and the oxidation of metalloporphyrins is probably reasonably fast at all pH. Reaction (2) is probably the rate limiting step when using mCPBA as oxidant and the overall reaction is faster at lower pH. The optimal pH for the oxidation of veratryl alcohol by a i 6TSPPFeO/t-BuOOH and TDCSPPFeCl/t-BuOOH was pH 5 and pH 6, respectively. Kinetic data for both reaction (1) and (2) are needed to give a detailed explanation of these observations. It is obvious by exarnining Table 2.1 that when using the same oxidant, it is 108 normally true that a manganese porphyrin has a higher optimal pH than its corresponding iron porphyrin (TDCSPPMnCl compared to TDCSPPFeCl and Cl16TSPPMnCl compared to Cl16TSPPFeCl) and a hexadecachloroporphyrin complex has the same or higher optimal pH compared to its corresponding octachloroporphyrin complex (Cl16TSPPFeCl over TDCSPPFeCl and Cl16TSPPMnCl over TDCSPPMnCl). This can be explained as follows. A metalloporphyrin is oxidized more readily at higher pH since its oxidation potential decreases with increasing pH2-25 and it is known that a manganese porphyrin has a higher oxidation potential than its iron anolog26 and a hexadecachloroporphyrin complex (Cl16TSPPFeCl or Cl16TSPPMnCl) is more electron deficient than a octachloroporphyrin complex (TDCSPPFeCl or TDCSPPMnCl).27 A higher pH will facilitate the oxidation of the metalloporphyrins with higher oxidation potentials. 2.1.3.2 Oxidant Hydrogen peroxide, t-BuOOH, /wCPBA, and NaCIO have all been used as oxidants to oxidize various metaUoporphyrins. The two-electron oxidation of metalloporphyrins by acyl- or alkylhydroperoxides (ROOH, R=acyl, alkyl, or H) involves the heterolytic O-O bond scission and the reaction is therefore accelerated by decreased electron density of R. In other words, the reaction of a metaUoporphyrin and ROOH is facilitated by a decrease in the pKa value of ROH.28"33 Among the peroxide type of oxidants used in this study, mCPBA should give the fastest reaction since the pKa order is mCBA < FLO < t-BuOH. It was observed that under the same conditions, the oxidation of veratryl alcohol by mCPBA was faster than that by t-BuOOH and r L 0 2 when the iron porphyrins (TDCSPPFeCl, Cl16TSPPFeCl) were the catalysts. This is also true at low pH when the 109 manganese porphyrins (TDCSPPMnCl, Cl 1 6 TSPPMnCl) were used as catalysts. For TDCSPPMnCl or Cl 1 6 TSPPMnCl catalyzed oxidation of veratryl alcohol at high pH (>8), however, the rate of reaction is much faster with hydrogen peroxide than with /nCPBA. This is probably because the manganese porphyrins were oxidized by mCPBA and hydrogen peroxide through different pathways. The oxidation of TDCSPPMnCl and Cl 1 6 TSPPMnCl by hydrogen peroxide may involve homolylic cleavage34,35 of the O-O bond of hydrogen peroxide. The oxidation of veratryl alcohol by t-BuOOH was efficient only when the iron porphyrins, TDCSPPFeCl and Cl 1 6TSPPFeCl, were used as catalysts. TDCSPPMnCl or Cl 1 6 TSPPMnCl was unable to oxidize veratryl alcohol using t-BuOOH as oxidant. Hydrogen peroxide caused a rapid destruction of Cli 6TSPPFeCl and therefore was not a suitable oxidant for this catalyst. The oxidation of veratryl alcohol by hydrogen peroxide was favoured at low pH when TDCSPPFeCl was the catalyst. When the manganese porphyrins TDCSPPMnCl or Cl 1 6 TSPPMnCl were the catalysts, however, the reaction was much faster at high pH as shown in Figure 2.34 and Figure 2.38. Hydrogen peroxide is a cheap, clean oxidant and would be the first choice if biomimetic catalysts are to be used in industry. The manganese porphyrins TDCSPPMnCl and Cl 1 6 TSPPMnCl seem to be better catalysts than the iron porphyrins for using hydrogen peroxide as oxidant. The optimal pH of these reactions, unfortunately, is high and as described above, the oxidant is not used efficiently at high pH due to the formation of superoxide. The optimal pH of lignin peroxidase is around pH 3 s 6 (varies for different isoenzymes). These manganese porphyrin/hydrogen peroxide systems may not mimic the lignin peroxidase at high pH (pH 10). On the other hand, effective degradation of lignin by lignin peroxidase at pH 3 has not been reported and the pH dependency of lignin degradation and pulp bleaching by lignin peroxidase or 110 biomimetic catalysts has not been studied, pH 3 is not necessarily the best pH for lignin oxidation or pulp bleaching. The manganese porphyrin/hydrogen peroxide systems, therefore, may have an advantage by working at high pH where the lignin peroxidase had no activity. The metalloporphyrins can also use NaCIO as oxidant for the oxidation of veratryl alcohol. The optimal pH for the reaction was pH 10 when the hexadecachloroporphyrin complexes (Cli 6TSPPMnCl, Cl 1 6TSPPFeCl) were the catalysts. The reaction was favoured at around neutral pH when the octachloroporphyrin complexes (TDCSPPFeCl, TDCSPPMnCl) were the catalysts. Sodium hypochlorite is also a possible oxidant suitable for industrial applications. It is important to find out if this oxidant will lead to the formation of any chlorinated organic compounds when used in the pulp and paper industry, since elimination of chlorinated pollutants is the major goal of using biotechnology in the pulp and paper industry. 2.1.3.3 The presence of manganese complexes As discussed earlier, superoxide is formed in the oxidation of veratryl alcohol. Under acidic conditions, superoxide dismutates rapidly giving oxygen and hydrogen peroxide and the reaction can be facilitated by the formation of additional hydrogen peroxide. At high pH, however, superoxide is relatively stable and reacts with the metalloporphyrin which decreases its catalytic activity as shown in Scheme 2.3 using Cl 1 6 TSPPMnCl as an example. In the presence of superoxide, reaction B competes with reaction A for the manganese(III) porphyrin catalyst and the former reaction leads to the formation of the catalytically inactive species Cl 1 6TSPPMn(II). Though the inactive manganese(II) porphyrin can be converted back to the manganese(III) porphyrin 111 C I 1 6 T S P P M n v = 0 Scheme 2.3 The effect of superoxide on the catalytic cycle of Cl 1 6 TSPPMnCl during the oxidation of veratryl alcohol, V A : veratryl alcohol; (VA) + : veratryl alcohol cation radical; [O]: oxidant (reaction pathway C), oxidant is required for this process and this reaction thus competes with the catalytic reaction pathway A for oxidant Both the rate and yield of the oxidation of veratryl alcohol is decreased due to the formation of superoxide. Manganese (U) complexes have been used as models for superoxide dismutase.21,22 The presence of manganese(U)-lactate increased the activity of both lignin peroxidase and the manganese(II)-dependent peroxidase.18 It was expected that the presence of manganese(JI)-complexes could also facilitate the oxidation of veratryl alcohol by metaUoporphyrins at high pH. Manganese(U)-lactate which was found to be able to stimulate the activity of the manganese(n)-dependent peroxidase was used as the manganese(II)-complex in this study. Since the mixture of manganese(n) sulfate and lactic acid (0.1 mM and 6 mM concentration, respectively) was not stable at high pH due to the formation of manganese(H) hydroxide, the effect of manganese(U)-lactate on the oxidation of veratryl alcohol was examined only at pH 5 and pH 6. As shown in Figures 2.40 and 2.41, the presence of manganese(IT)-lactate increased the rate of oxidation of veratryl alcohol by TDCSPPFeClAwCPBA or Cl16TSPPMnCI/mCPBA. It was noticed that in the presence of manganese(TI)-lactate, the absorption at 310 nm (Xta„ of veratraldehyde) started decreasing after reaching a maximum. This was due to the interference of manganese(IJI)-lactate, which also absorbed at 310 nm. Manganese(DT)-lactate decomposed slowly in aqueous solution. The stable absorbance at 310 nm 24 hours after the reaction was still higher than that of the control experiment without nianganese(TJ)-lactate, mdicating that the yield of veratraldehyde was indeed increased by the presence of manganese(n)-complexes. The observation that manganese(ni)-lactate was formed and accumulated during the reaction also suggested that manganese(n)-lactate was not a good model for superoxide dismutase in this case. Manganese(n) complexes can also be oxidized by high valent 113 0.0 1.0 2.0 3.0 Time (min) Figure 2.40 The effect of manganese(II)-lactate (4xl0' 9 mole and 2.4xl0' 7 mole, respectively) on the oxidation of veratryl alcohol (5x10"* mole) by TDCSPPFeCl (1x10* mole) and mCPBA (2x10* mole) at room temperature in 3 mL 0.1 M citrate buffer, 1, pH 5 in the presence of M n 2 + ; 2, pH 6 in the presence of Mn 2 + ; 3, pH 5 in the absence of M n 2 + ; 4, pH 6 in the absence of M n 2 + 114 A 2.00 -0.0 1.0 2.0 3.0 Time (min.) Figure 2.41 The effect of manganese(II)-lactate (4x l0 9 mole and 2.4xl0"7 mole, respectively) on the oxidation of veratryl alcohol (5x10"* mole) by Cl 1 6 TSPPMnCl ( l x l O 4 mole) and mCPBA (2x10* mole) at room temperature in 3 mL 0.1 M citrate buffer, 1, pH 5 in the presence of M n 2 + ; 2, pH 5 in the absence of M n 2 + ; 3, pH 6 in the presence of Mn 2 + ; 4, pH 6 in the absence of M n 2 + 115 metaUoporphyrins to manganese(ITI) complexes in addition to their reaction with superoxide. The manganese(II)-lactate, therefore, competed with the veratryl alcohol substrate for the high valent metaUoporphyrins. This side reaction, however, might be minimized by using a ligand other than lactate or by changing the concentration of Mn(H). No effort was made to search for a better superoxide dismutase model than manganese(n)-lactate. It should be added that the formation and accumulation of manganese(TJLI)-lactate is not necessarily a disadvantage to the system. The manganese(II)-dependent peroxidase uses manganese(Ti) complexes as mediators for the oxidation of its substrates. Although manganese(III) complexes such as Mn(IU)-lactate are unable to oxidize veratryl alcohol, they can oxidize MnP substrates such as polymer dyes and phenolic compounds. Non-phenolic lignin model compounds can also be oxidized under certain conditions.37 Further study is needed to understand the role of manganese(II) and MnP in the biodegradation of lignin. 2.1.3.4 The effect of imidazole Peroxidases, including lignin peroxidase38,39 and Mn(U)-dependent peroxidase40, have an iron protoporphyrin IX with a proximal-bound imidazole group of Wstidine at their active sites. Imidazole has been used to facilitate metaUoporphyrin catalyzed reactions.41"42 The coordination of imidazole to the metal increases its electron density and thus aids the heterolytic 0-0 cleavage of bound peroxides. The epoxidation of alkenes in organic solvents by meso-tetraphenylporphyrin iron chloride (TPPFeCl) or meso-tetraphenylporphyrin manganese chloride (TPPMnCl) using either hydrogen peroxide or alkyl hydrogen peroxides as oxidant was successful only in the presence of 116 imidazole.4M2 It has been reported that manganese porphyrins ligate only one imidazole per molecule43 and in methylene chloride the rate- of oxidation of TPPMnCl by alkyl hydroperoxides or percarboxyUc acids was increased by 10M04 fold in the presence of imidazole.30 In aqueous solution, however, the rate of the oxidation of manganese porphyrin 25 by hydrogen peroxide was increased by only about 10 fold by adding imidazole to the reaction mixture.26 Metalloporphyrins with attached imidazole groups have also been used as peroxidase models.33/4 While coordination of one imidazole group to an iron porphyrin increased its oxidation rate, attaching an additional imidazole group to the porphyrin inhibited its oxidation.44 The oxidation of veratryl alcohol (1x10s mole) in the presence of imidazole (0-6x10* mole) by various metalloporphyrin (TDCSPPFeCl, Cl16TSPPFeCl, TDCSPPMnCl, or Cl16TSPPMnCl, 3x10* mole) and oxidant (mCPBA, NaCIO, H202. or t-BuOOH, 5x10* mole) combinations were studied. The presence of imidazole had very little effect on the rate of veratryl alcohol oxidation in most cases and the reactions were even inhibited by the presence of a large excess of imidazole. The only example where reasonable acceleration was observed was the oxidation of veratryl alcohol by Cl16TSPPMnCl and H202 as shown in Figures 2.42 and 2.43. The very small stimulating effect of imidazole is probably due to the use of aqueous solution and the fact that one to one coordination of imidazole to the metaUoporphyrin is difficult to achieve.26,30 117 0.0 1.0 2.0 3.0 4.0 5.0 6.0 Time (min.) Figure 2.42 The effect of imidazole on the oxidation of veratryl alcohol ( l x l O 5 mole) by hydrogen peroxide (5x10-* mole) and Cl 1 6 TSPPMnCl (3x10"* mole) at room temperature in 3 mL 0.1 M pH 9.2 borate buffer, the amount of imidazole (mole): 1, 0; 2, 1.5X10"6; 3, 3x10"*; 4, 6x10-* 118 0.0 2.0 4.0 6.0 8.0 Time (min.) Figure 2.43 The effect of imidazole on the oxidation of veratryl alcohol ( l x l O 5 mole) by hydrogen peroxide (5x10* mole) and Cl 1 6 TSPPMnCl (3X10-8 mole) at room temperature in 3 mL 0.1 M pH 10 phosphate buffer, the amount of imidazole (mole): 1, 0; 2, 3x10 s; 3, 6x10* 119 22 Conclusion The stability of the metal complexes of TPP derivatives towards excess oxidants is dramatically increased by introducing chlorine at the phenyl ring and Impositions. The relative and absolute stability of the metalloporphyrins also depends on pH, the type of oxidant being used, and the presence or absence of substrates. Two oxidized intermediates, probably oxomanganese(IV) and oxomanganese(V) porphyrins, were observed spectrally when TDCSPPMnCl or Cl16TSPPMnCl were oxidized. Only one oxidized intermediate of the iron porphyrins (TDCSPPFeCl or Cl,6TSPPFeCl) was observed and its nature is not clear. Cl16TSPPMn(D.) and Cl16TSPPFe(DJ)-02\ probably formed by the reaction of the metal(IU) porphyrin and superoxide, were observed in the oxidation of the respective metaUoporphyrin in the presence of veratryl alcohol. The catalytic activity of the metalloporphyrins for the oxidation of veratryl alcohol depends on the oxidant being used, the pH of the solvent, and to a lesser extent, on the presence of manganese(II)-complexes and imidazole. The largest catalytic activity for the metalloporphyrins was observed at low pH using mCPBA as oxidant. At high pH (pH 10), TDCSPPMnCl exhibits the largest catalytic activity using hydrogen peroxide as oxidant. The optimal pH for maximal catalysis of a metaUoporphyrin/oxidant combination depends on both the type of catalyst and oxidant. TDCSPPMnCl and Cl16TSPPMnCl usually have a higher optimal pH than their conesponding iron porphyrins (TDCSPPFeCl and Cl16TSPPFeCl) and the rctacMoroporphyrins (TDCSPPFeCl and TDCSPPMnCl) usually have a lower optimal pH than their corresponding hexadecacMoroporphyrins (Cl,6TSPPFeCl and Cli6TSPPMnCl) when using the same oxidant. 120 The oxidation of veratryl alcohol is stimulated by increasing buffer concentration or by adding some inorganic salts, supporting a cation radical mechanism of the reaction. The presence of high concentrations of sodium fluoride or sodium nitrate, however, was found to inhibit the reaction completely. Oxidized intermediates of iron and manganese porphyrins have been observed when they are oxidized at low temperature in organic solvents.3-10 The observation of oxidized intermediates of TDCSPPMnCl and Cl 1 6 TSPPMnCl by UV-vis spectroscopy at room temperature is interesting and needs further study. Techniques such as ESR, NMR, IR, Resonance Raman spectroscopy and magnetic measurements are needed to characterize these intermediates. Only one oxidized intermediate of TDCSPPFeCl and Cl 1 6 TSPPFeCl was observed, the use of stop-flow apparatus may enable the detection of the second oxidized intermediate. The formation of Cl 1 6 TSPPFe n -0 2 at room temperature indicates that it is stable and might be a good model for haemoglobin. 2.3 Experimental 2.3.1 General Equipment. A Hewlett Packard 8452A Diode Array UV-vis spectrophotometer was used for all the UV-vis spectroscopic measurements. A HP 5890A Gas Chromatograph equipped with a HP 17 capillary column (25 meters, 0.32 millimetre ID) and a FID (flame ionization detector) detector was used for all the G C analyses. Chemicals. Veratryl alcohol, purchased from Aldrich, was vacuum distilled before use. Purified grade mCPBA from Fisher Scientific Company was used in this experiment. Mixed strong acidic/strong basic ion exchange resin Amberlite MB-3 and 121 strong acidic ion exchange resin Amberlite IR-120 were purchased from Mallinckrodt Corporation. Strong basic ion exchange resin Rexyn® was from Fisher Scientific Company. A l l the ion exchange resins were washed with 5 volumes of deionized water on the column before use. Other chemicals were of reagent grade and used without further purification. The phosphate buffers were made from appropriate amounts of phosphoric acid, K H 2 P 0 4 , K 2 H P 0 4 , and/or K 3 P 0 4 depending on the pH of the respective buffer. The final pH of the buffers were adjusted to the desired value using dilute NaOH or H 3 P0 4 . Citric acid was neutralized with sodium hydroxide solution to the desired pH (pH 5 or 6) and then diluted with deionized water to give a final concentration of 0.1 M . Borate buffer (pH 9.2) was made by diluting the commercially available powder from Fisher Scientific Company. A l l the metalloporphyrins were synthesized following a modified procedure of Nakano et a/.45 Meso-tetra(2,6-cuchlorophenyl)porphyrin and me^o-tetra(2,6-dichlorophenyl)-6-octachloroporphyrin, provided by Dr. Tilak Wijesekera of this laboratory were first sulfonated using fuming sulfuric acid to give we5o-tetra(2,6-dichloro-3-sulfonatophenyl)porphyrin and mes<>tetra(2,6-cuchloro-3-sulfonatophenyl)-J3-octachloroporphyrin, respectively. The sulfonated porphyrins were then metallated in D M F to give the metalloporphyrins 12-15. Me5o-tetra(2,6-dichlorophenyl)porphyrin (300 mg) was suspended in 14 mL of fuming sulfuric acid and heated with stirring at 165 °C for 7 hours. The reaction mixture was then cooled, added slowly to 50 mL ice cooled water, and filtered. The filtrate was further diluted with 75 mL of water and neutralized with NaHC0 3 (65 gram in 100 mL water). Ethanol (250 mL) was added to the solution to precipitate the sodium sulfate, which was removed by filtration. Sodium sulfate was precipitated once 122 more by adding 500 mL ethanol to the clear filtrate. The solution containing the product, sodium sulfate, and possibly small amount of sodium bicarbonate, was evaporated on a Rotavapor. The residue was dissolved in 10 mL deionized water and filtered. Remaining sodium bicarbonate in the filtrate was neutralized carefully with 10% HC1 to neutral pH. The solution was passed slowly through a glass column (3x30 cm) packed with mixed strong acidic and strong basic ion exchange resin (Amberlite MB-3, Mallinckrodt Inc.). The eluent was dried to give a fine black microcrystalline solid. Me5o-tetra(2,6-dichlorophenyl)-6-octachloroporphyrin was sulfonated similarly. The sulfonated porphyrins were metallated following reported procedures.45 The purification of the metalloporphyrins45 was considered inappropriate. The procedure was modified by using ion exchange resins. The crude product obtained via precipitation using acetone was found difficult to dissolve in water due to the presence of iron oxides. Dilute HC1 was added to completely dissolve the residue in 10 mL of deionized water. The solution was then passed through a 3x30 cm ion exchange resin (strong acidic resin, Amberlite IR-120) column to remove all the free iron in the solution. Evaporating the eluent gave a fine black/blue microcrystalline powder. Ferric ions were found to be retained efficiently on the column and the metalloporphyrin was therefore uncontaminated. After washing the column with saturated sodium chloride, addition of potassium thiocyanate to the yellow eluent gave a deep red color, indicating the presence of iron. When the metalloporphyrin solution after ion exchange was passed through a small column packed with fresh ion exchange resin, no iron was retained on the column. The synthesis of TDCSPPFeOH. TDCSPPFeCl (10 mg) was dissolved in 1 mL of water and dilute sodium hydroxide was added until the iron porphyrin showed a 123 sharp single Soret band in its UV-vis spectrum. The solution was then passed through an ion exchange column (1.5x15 cm) packed with strong basic ion exchange resin (Rexyn* 201, Fisher Scientific Company). The eluent was evaporated on a Rotavapor. 2.3.2 The stability of the metalloporphyrins towards excess oxidants The stability of the metalloporphyrins towards excess hydrogen peroxide at pH 3 in the presence of veratryl alcohol was determined as follows. Hydrogen peroxide (3x10* mole) was added to a solution (4 mL 0.1 M phosphate buffer, pH 3) containing the porphyrin (3x10* M) and veratryl alcohol (1x10* M) at room temperature. The Soret absorption changes of the metalloporphyrin with time were followed by UV-vis spectroscopy. At pH 10, addition of hydrogen peroxide to the manganese porphyrins (TDCSPPMnCl, Cl16TSPPMnCl) led to the formation of relatively stable oxidized intermediates. The extinction coefficients of these oxidized intermediates were not easy to detemiine and the quantification of the total concentration of the various porphyrin species was thus difficult. This problem was solved by reducing the various oxidation states of the manganese porphyrins to Mn(TJ) porphyrin by adding excess sodium borohydride (1 mg) immediately before the spectral measurement. The stability of the various metalloporphyrins was compared by plotting (A^/As^Jo against time t, where (ASaJt is the Soret absorbance of the Mn(H) porphyrin produced by reducing the reaction rnixture with excess NaBIL, at time t, and ( A J ^ Q is the Soret absorbance of the Mn(II) porphyrin after treating the reaction mixture with NaBIL, at time 0. Hydrogen peroxide (1.5x10* mole) was added to 20 mL 0.1 M pH 10 phosphate buffer containing 1.5x1a7 mole of TDCSPPMnCl (or Cl16TSPPMnCl) and 5x10s mole of 124 veratryl alcohol. 3 mL of the reaction mixture was taken out at time 0, 2, 4, 6, 8 and 10 minutes, reduced with NaBFL, (1 mg) and subject to UV-vis measurement. The stability of TDCSPPFeCl and Cl 1 6 TSPPFeCl was determined similarly as for the experiments at pH 3. , The stability of TDCSPPFeCl towards excess mCPBA (or H A ) at various pH in the absence of veratryl alcohol was also determined by UV-vis spectroscopy. mCPBA (3xl0' 7 mole dissolved in 10 | i L acetonitrile) was added to a solution (3 mL 0.1 M phosphate buffer) containing 3x10* mole of TDCSPPFeCl and the change of the Soret absorption of TDCSPPFeCl with time was recorded. The results were plotted as ( A s o J A A s c J o against time t. 2.3.3 The oxidized intermediates of the metalloporphyrins A l l the reactions were carried out at room temperature. The oxidation of the metalloporphyrins by various oxidants in the presence or absence of veratryl alcohol was carried out in a quartz UV-vis cell containing 3 mL solution. The metalloporphyrin (1 piiole/mL in water) and veratryl alcohol (1 M solution in water) were added. The oxidants were dissolved in appropriate solvents (mCPBA and t-BuOOH in acetonitrile, H 2 0 2 and NaCIO in water, 0.1 M in concentration) and added using a syringe. The reactions were followed by UV-vis spectroscopy. The oxidation of ClJ6TSPPMn(II) by NaCIO. When NaCIO QxlO" 7 mole) was added to a solution of Cl 1 6TSPPMn(II) generated by adding 2x10 s mole of NaCIO to a solution (3 mL 0.1 M phosphate buffer, pH 7) containing Cl 1 5 TSPPMnCl (3xl0' 8 mole) and veratryl alcohol (1x10 s mole), a small but very rapid decrease in the Soret band of the Mn(U) porphyrin was observed. The peak increased again in intensity afterwards. 125 Addition of 5x10* mole more NaCIO converted the spectrum instantly to that of the Mn(DI) porphyrin. The oxidation of Cl16TSPPMnCl by NaCIO in the presence of manganese(II)-lactate. The reaction mixture contained in 3 mL 0.1 M citrate buffer (pH 5 or 6) 3x10* mole of Cl16TSPPMnCl and 1x10s mole of veratryl alcohol. Manganese(H) sulphate and (L)-lactic acid were added to a final concentration of ImM and 60mM, respectively. 2x10s mole of NaCIO was added to start the reaction. The oxidation of Cl16TSPPFeCl by NaCIO in the presence of manganese(II)-lactate was carried out under similar conditions. The oxidation of Cl16TSPPFe(III)-02- by NaCIO was similar to the oxidation of Cl16TSPPMn(TI). 23.4 Factors affecting the catalytic activity of the metalloporphyrins All the reactions were carried out at room temperature. The oxidation of veratryl alcohol by mCPBA catalyzed by various metalloporphyrins. 2x10* mole of mCPBA dissolved in 20 pX acetonitrile was added to 4 mL solution (0.1 M phosphate or citrate buffer) containing 1x10* mole of metaUoporphyrin and 5x10* mole of veratryl alcohol. The formation of veratraldehyde was followed by UV-vis spectroscopy at 310 nm. The oxidation of veratryl alcohol by various metalloporphyrins using NaCIO, t-BuOOH, or H202 as oxidants. The reaction mixture (4 mL 0.1 M phosphate, citrate or borate buffer) contained 3x10* mole of metalloporphyrin, 5x10* mole of oxidant and 1x10s mole of veratryl alcohol. The oxidation of veratryl alcohol in the presence of manganese(II)-lactate 126 (Figures 2.40, 2.41). mCPBA (2x10* mole) was added to a solution containing 1x10* mole of metalloporphyrin (TDCSPPFeCl or Cl 1 6TSPPFeCl), 5x10* mole of veratryl alcohol, 4xl0 ' 9 mole of manganese(Tf) sulphate and 2.4xl0' 7 mole of lactic acid. The formation of veratraldehyde was followed by UV-vis spectroscopy at 310 nm. The oxidation of veratryl alcohol in the presence of imidazole. The conditions for all the reactions were similar. A typical reaction mixture contained in 3 mL solvents 3x10* mole of metalloporphyrin, 5x10* mole of oxidant (H 2 0 2 and NaCIO were added as aqueous solution, t-BuOOH and mCPBA were added as acetonitrile solution), 1x10* mole of veratryl alcohol and various amounts of imidazole, ranging from 2 to 200 times excess (molar ratio, imidazole over metalloporphyrin), as specified in Figures 2.42 and 2.43. The formation of veratraldehyde was followed by UV-vis spectroscopy at 310 nm. 127 References for Chapter 2 1. T.G. Traylor, and S. Tsuchiya, Inorg. Chem. 26, 1338, 1987. 2. N . Carnieri, A . Haniman, and G. Porter, J. Chem. Soc. Dalton Trans. 931, 1982. 3. J.T. Groves, R.T. Haushalter, M . Nakamura, T.E. Memo, and B.J. Evans, J. Am. Chem. Soc. 103, 2884, 1981. 4. J.T. Groves, T.E. Memo, and R.S. Meyers, J. Am. Chem. Soc. 101, 1032, 1979. 5. D. Chin, J. Del Gaudio, G.N. Lamar, and A . L . Balch, J. Am. Chem. Soc. 99, 5486, 1977. 6. J.T. Groves, and Y . Watanabe, J. Am. Chem. Soc. 110, 8443, 1988. 7. J.T. Groves, and M . K . Stem, J. Am. Chem. Soc. 110, 8628, 1988. 8. J.T. Groves, and M . K . Stem, J. Am. Chem. Soc. 109, 3812, 1987. 9. R.S. Czernuszewicz, Y.O. Su, M . K . Stem, K . A . Macor, D. Kim, J.T. Groves, and T.G. Spiro, J. Am. Chem. Soc. 110, 4158, 1988. 10. M . Schappacher, and R. Weiss, Inorg. Chem. 26, 1190, 1987. 11. B.C. Schardt, F.J. Hollander, and C.L. Hi l l , J. Chem. Soc., Chem. Commun. 765, 1981. 12. J.A. Smegal, and C.L. Hi l l , J. Am. Chem. Soc. 105, 3515, 1983. 13. J.T. Groves, W.J. Kruper, and Jr. R.C. Hanshalter, J. Am. Chem. Soc. 102, 6375, 1980. 14. J.T. Groves, and Y . Watanabe, Inorg. Chem. 25, 4808, 1986. 15. J.T. Groves, Y . Watanabe, and T.J. McMurry, J. Am. Chem. Soc. 105, 4489, 1983. 128 16. S.E. Creager, S.A. Raybuck, and R.W. Murray, J. Am. Chem. Soc. 108, 4225, 1986. 17. N . Carnieri, A . Harriman, and G. Porter, J. Chem. Soc., Dalton Trans. 1231, 1982. 18. J.J. Bono, P. Goulas, N . Portet, and J.L. Sens, Collq INRA 40 (Lignin Enzymic Microb. Degrad.), 87, 1987. 19. J.S. Valentine, and A .E . Quinn, Inorg. Chem. 15, 1997, 1976. 20. B . M . Hoffman, C S . Weschler, and F. Basolo, J. Am. Chem. Soc. 98, 5473, 1976. 21. B. Halliwell, NeuroToxicology, 5, 113, 1984. 22. J. Epp, S. Fairchild, G. Erickson, and W.H. Koppenol, in Superoxide and Superoxide Dismutase in Chemistry, Biology, and Medicine. Ed. G. Rotilio, Elsevier Science Pub. 1986, p76. 23. Cynthia D. Millis, unpublished results. 24. M . Tien, T.K. Kirk, C. Bull , and J.A. Fee, J. Biol. Chem. 261, 1687, 1986. 25. M.F. Zipplies, W.A. Lee, and T.C. Bruice, J. Am. Chem. Soc. 108, 4433, 1986. 26. P.N. Balasubramanian, E.S. Schmidt, and T.C. Bruice, J. Am. Chem. Soc. 109, 7865, 1987. 27. T. Wijesekera, A . Matsumoto, D. Dolphin, and D. Lexa, Angew. Chem. 1990, in press. 28 L . -C . Yuan, and T.C. Bruice, J. Am. Chem. Soc. 107, 512, 1985. 29. W.A. Lee, and T.C. Bruice, J. Am. Chem. Soc. 107, 513, 1985. 30. L . -C . Yuan, and T.C. Bruice, J. Am. Chem. Soc. 108, 1643, 1986. 31. W.A. Lee, L . -C . Yuan, and T.C. Bruice, J. Am. Chem. Soc. 110, 4277, 1988. 32. T.C. Bruice, P.N. Balasubramanian, R.W. Lee, and J.R.L. Smith, J. Am. Chem. 129 Soc. 110, 7890, 1988. 33. T.G. Traylor, and J.P. Ciccone, J. Am. Chem. Soc. I l l , 8413, 1989. 34. T.C. Bruce, Aldrichimica Acta, 21, 87, 1988. 35. J.R.L. Smith, P.N. Balasubramanian, and T.C. Bruice, J. Am. Chem. Soc. 110, 7411, 1988. 36. N . Kirkpatrick, and J .M. Palmer, Appl. Microbiol. Biotechnol. 30, 305, 1989 37. M . H . Gold, H . Wariishi, K . Valli , M . B . Mayfield, V.J . Nipper, and D. Probnow, Biotechnol. in the Pulp and Paper Industry (Proceedings of the 4th International conference), Raleigh, NC, USA, May 16-19, 1989, in press. 38. D. Kuila, M . Tien, J.A. Fee, and M.R. Ondrias, Biochem. 24, 3394, 1985. 39. L . A . Anderson, V . Renganathan, A . A . Chiu, T .M. Lohr, and M . H . Gold, J. Biol. Chem. 260, 6080, 1985. 40. Y . Mino, H . Wariishi, N.J. Blackburn, T . M . Lohr, and M . H . Gold, J. Biol. Chem. 263, 7029, 1988. 41. J.-P. Renaud, P. Battioni, J.F. Bartoli, and D. Mansuy, J. Chem. Soc, Chem. Commun. 888, 1985. 42. D. Mansuy, P. Battioni, and J.-P. Renaud, J. Chem. Soc., Chem. Commun. 1255, 1984. 43. S. Neya, I. Morishima, and T. Yonezawa, Biochem. 20, 2610, 1981. 44. T.G. Traylor, W.A. Lee, and D.V. Stynes, J. Am. Chem. Soc. 106, 755, 1984. 45. T. Nakano, T.P. Wijesekera, D. Dolphin, T.E. Maione, T.K. Kirk, and R.L. Farrell, U.S. patent 4,892,241, 1989. 130 Chapter 3 Oxidation of lignin model compounds catalyzed by meso-tetra(2,6-dichloro-3-suIfonatophenyl)porphyrin iron chloride (TDCSPPFeCl) S03H SO„H Figure 3.1 The structure of TDCSPPFeCl 3.1. Results and discussion Due to the complex structure of lignin, model compounds representing separate portions of lignin have been widely used in lignin biodegradation studies. Though it is not clear if lignin and lignin model compounds are degraded by Ugnin degrading enzymes through the same pathway, Ugnin model compounds have played an important role in suggesting the mechanism of Ugnin degradation. In this study, Ugnin model compounds were used to study the potential of the metaUoporphyrins to degrade various 131 substructures of lignin. 3.1.1. The oxidation of veratryl alcohol and veratryl acetate Veratryl alcohol is a secondary metabolite of Phanerochaete chrysosporium1* and plays a unique role in this process.3 It was reported that veratryl alcohol could stimulate lignin degradation4,5 and benzo(a)pyrene6 oxidation by lignin peroxidase by protecting the enzyme from inactivation7'9 in addition to functioning as a mediator at the same time (see chapter 4 of this thesis). Veratryl alcohol can also induce the biosynthesis of lignin peroxidase by P. chrysosporium}0 In addition, veratryl alcohol is also a convenient lignin model compound It is readily oxidized to give veratraldehyde by Ugnin peroxidase in the presence of hydrogen peroxide. On the other hand, veratraldehyde is readily reduced back to veratryl alcohol which accumulates in the P. chrysosporium culture.2 This redox cycle of veratryl alcohol seemed to be energetically wasteful as the oxidizing power is used to oxidize veratryl alcohol instead of lignin. Leisola et al11 first found that veratryl alcohol could be oxidized to give aromatic ring cleavage products. Later studies12 showed that veratryl alcohol was rapidly metabolized to carbon dioxide by carbon limited cultures of P. chrysosporium, compared to the slow and incomplete degradation in nitrogen limited cultures. It was also shown12 that veratraldehyde, the major oxidation product of veratryl alcohol by lignin peroxidase, could not be metabolized directly by the fungus, instead, it was reduced back to veratryl alcohol in the culture, and metabolized to carbon dioxide via intermediating quinones and aromatic ring cleavage products. The degradation of veratryl alcohol by lignin peroxidase and biorrtimetic catalysts has been extensively studied11'16 and the products including two y-lactones, two 6-132 lactones and three quinones have been found (Figure 3.2). The proposed mechanism for the production of these compounds was that initial one-electron oxidation of veratryl alcohol gave a cation radical, attack of which by water gave a carbon-centred radical, coupling of which with molecular oxygen lead to aromatic ring cleavage (pathway A) or demethoxylation products (quinones) (pathway B). Quinones can also be formed through pathway C as shown in Figure 3.3. Oxidation of veratryl alcohol by TDCSPPFeCl/mCPBA at room temperature in pH 3 aqueous buffer gave veratraldehyde as the major product (>80%) along with small amounts of ring cleavage products 26, 28, 29, and quinone 30 (Figure 3.2). When 4-ethoxy-3-methoxybenzyl alcohol was oxidized under the same conditions, lactones 28 and 29 were also found as products, conforming their 5-lactone structure. The quinones 31 and 32 formed in the oxidation of veratryl alcohol by lignin peroxidase were not found in this study. As will be described later, however, quinone 32 was found as a product of the oxidation of veratryl alcohol by TDCSPPFeCl/mCPBA using methanol as solvent. When veratryl acetate was oxidized by TDCSPPFeCl/mCPBA in pH 3 aqueous buffer, quinone 34, 3,4-dimethoxyphenol 35, and a direct ring cleavage product 36 were isolated as the major products (Figure 3.4). Small amounts of veratraldehyde and veratryl acetate were also found. Quinones 31 and 32 (Figure 3.2) have been found as phenyl-alkyl cleavage products of veratryl alcohol.11,12,16 To our knowledge, this is the first time a phenol (35) has been found as a phenyl-alkyl cleavage product of a veratryl alcohol derivative. The quinone 34 found in this reaction is a analog of 30 found in the oxidation of veratryl alcohol, indicating that acetylation of veratryl alcohol to give veratryl acetate does not inhibit its demethoxylation. 133 Figure 3.2 The oxidation of veratryl alcohol by lignin peroxidase or biomimetic systems C H 2 O H _ C H 2 O H H 2 Q 0 C H 3 0 C H 3 C H 2 O H O ^ H C H 3 - 0 0 ^ 1-H 2 0 C H 2 O H 6 C 0 2 C H 3 C 0 2 C H 3 33 0 \ C H 3 0 C H 2 O H OH O C H 3 0 C H 3 02 C H 2 O H J C H 3 0 0 OH O C H 3 \ „ S 0 • 0 - 0 . C H 2 O H OH O C H 3 | 0 C H 3 1 B 02 C H 2 O H OH O C H 3 O C H 3 C H 2 O H 0 O C H 3 \ \ - CHbOH C H 2 O H 0 O C H 3 C H 2 O H H 2 0 H, HCK 0 O C H 3 C H 2 O H ' L ^ 0 C H 3 + 0 V 0 . o S > 0 C H 3 X X A 0 O C H 3 O C H 3 Figure 3.3 Proposed mechanisms for the oxidation of veratryl alcohol 135 Isolation of the direct ring cleavage product 36 supports the proposed mechanism of aromatic ring cleavage of veratryl alcohol.13 The aromatic ring cleavage of veratryl alcohol gave initially 33 (Figure 3.3), intermolecular transesterification gave the C H 2 O A C J ^ O C H 3 0 C H 3 Figure 3.4 The oxidation of veratryl acetate in pH 3 aqueous buffer by TDCSPPFeCl and mCPBA lactones shown in Figure 3.3. Compound 36 can be considered as a "trapped" analogue of intermediate 33. A mechanism for the formation of 36 is proposed in Figure 3.5. We learned after finishing this work that Schmidt et a/1 7 studied the degradation of veratryl methyl ether by lignin peroxidase and found a direct ring cleavage product similar to 36. Veratraldehyde was the major product of veratryl methyl ether oxidation.17 A Ca-ether bond, therefore, can be cleaved by lignin peroxidase, most probably through a one-electron oxidation mechanism.17,18 The formation of veratraldehyde from the oxidation of veratryl acetate, though in a small amount, C H 2 O A C + 3 5 C H O C H 2 O A C C 0 2 C H 3 C 0 2 C H 3 3 6 136 CH2OAc CH2OAc ^ CH2OAc I6L — ^ d l -Sr (&oh 0 C H 3 O C H 3 O C H 3 Y CH2OAc H n CH 2 OAc' CH2OAc C 0 ^ C ^ C H 3 - 0 * 0 ° - C H 3 C H 3 ^ 0 ° C ^ Figure 3.5 Possible mechanism for the formation of compound 36 suggests that the Ca-ester bond of lignin model compounds can also be cleaved by lignin peroxidase in a similar way. There have been reports that the oxidation reactions of lignin and Ugnin model compounds by enzymes19,20 or biomimetic catalysts21 vary in different solvents. As TDCSPPFeCl is soluble in both aqueous and polar organic solvents, the oxidation of veratryl alcohol and veratryl acetate in methanol was also studied to investigate possible solvent effects on Ugnin model compound oxidation. Oxidation of veratryl alcohol by TDCSPPFeCl/mCPBA in methanol gave veratraldehyde, methanol-incorporated products 37 and 38, and quinones 30 and 32 (Figure 3.6). The product distribution was dependent on the conditions used, when the 137 reaction was carried out open air with moderate stirring, veratraldehyde and 37 were produced in 47% and 42% yield, respectively. Other products were found only in very small amounts. Under highly aerobic condition (open air, vigorous stirring), the relative yield of veratraldehyde, 37, and 38 was 51%, 11%, and 32%, respectively. Under anaerobic condition, compound 38 was found only in trace amounts. CHO C H 2 O H 0 Figure 3.6 The oxidation of veratryl alcohol by TDCSPPFeCl/mCPBA in methanol. The structure of the methanol incorporated product 37 was confirmed as follows. Acetylation of 37 by acetic anhydride/pyridine gave 39. Oxidation of veratryl acetate by TDCSPPFeCl/mCPBA in methanol gave a product whose *H NMR and MS were the same as those of 39. Treating 37 with concentrated HC1 in aqueous THF gave compound 30 (Scheme 3.1). When veratryl alcohol was oxidized in ethanol, one solvent molecule was incorporated to give product 40. If 4-ethoxy-3-methoxybenzyl 138 CH 2OR-OCH 3 37 Rj^H, R2—R3—CH3 39 R 1 =COCH 3 , R 2=R 3=CH 3 40 R ^ H , R 2 =C 2 H 5 , R 3 =CH 3 41 R,=H, R 2=CH 3 , R,=CjH s 37 30 Scheme 3.1 The hydrolysis of compound 37. alcohol was used as the substrate, compound 41 instead of 37 was found as the product. The structure of 38 was elucidated by its 1 3 C N M R and by comparing its ' H C H 3 ° - C H ' ° C H 3 ^ n 38 R=CH 3 OCH3 4 2 R = C 2 H 5 OCH3 N M R and MS with those of 37. Oxidation of 4-ethoxy-3-methoxybenzyl alcohol in methanol gave product 42. A mechanism for the formation of 37 is proposed in Figure 3.7. Shimada et a/14 proposed two possible pathways for aromatic ring cleavage of veratryl alcohol and suggested that the one in which water (or hydroxide) specifically attacked the Cj-139 CH 2OH Figure 3.7 Possible mechanism for the formation of compound 37 position of a substrate cation radical was more plausible. The specific incorporation of methanol into the Q-position of veratryl alcohol to give compound 37 clearly indicates that the solvent methanol preferentially attacks the Cj-position of the veratryl alcohol cation radical. The formation of 37 also suggests that quinone 30 resulted from attack of water at the Q-position of the veratryl alcohol cation radical. Oxidation of veratryl acetate in methanol gave the methanol incorporated product 39 as a major product (82%). Small amounts of veratraldehyde and veratryl acetate were also found. It was interesting to observe that when veratryl alcohol was oxidized in ethanol, only one solvent molecule was incorporated to give compound 40 and that the 3-methoxy group of 40 did not exchange with the solvent. Aromatic ring cleavage products were not found when veratryl alcohol or veratryl acetate were oxidized in methanol, clearly suggesting the presence of solvent effect for the oxidation of lignin model compounds. 3.1.2. Oxidation of phenylpropane model compounds 43 141 Alkoxy substituted phenylpropane substructure (43) has been suggested to be present at the terminal chains of lignin.22 Tien and Kirk23 first found that lignin peroxidase could catalyze hydroxylation of the benzylic methylene group (Ca-hydroxylation) of 2-(3,4-(lmiethoxyphenyl)ethanol to give 3,4>dimethoxyphenylglycol. Ca-hydroxylation of l-(4-ethoxy-3-methoxyphenyl)propane by lignin peroxidase was also reported by Renganathan et al.24*5 Isotopic labelling experiments25 using 1802 or F y o showed that the oxygen of the Ca-hydroxy group was from molecular oxygen under aerobic conditions but came from water under anaerobic conditions, indicating a cation radical mechanism for the reaction. In addition to the Ca-hydroxylation product 47 reported earlier,24,25 oxidation of l-(4-ethoxy-3-methoxyphenyl)propane (44) by TDCSPPFeCl and t-BuOOH at room temperature in aqueous acetonitrile (pH 3) gave the direct ring cleavage product 45, quinone 46, and 48 as major products (Figure 3.8). The formation of the direct ring cleavage product 45 from 44 and the formation of ring cleavage product 36 from the oxidation of veratryl acetate support each other and extend our knowledge about the ring cleavage of lignin model compounds. Quinones analogous to 46 have been found in the oxidation of veratryl alcohol and veratryl acetate as described above. The formation of 48 is interesting and no similar products have been previously found from the oxidation of lignin model compounds. A mechanism for the alkyl-phenyl cleavage of phenolic model compounds has been proposed26 as shown in Figure 3.9. The formation of 50 resulted from a one-electron oxidation process and the alkyl-phenyl cleavage of 50 needed the presence of an acidic proton. In the case of 44, it has only an unreactive propyl side chain, hence alkyl-phenyl cleavage is unable to occur and product 48 can be considered as an analogue of the intermediate 50. A possible mechanism for the formation of 48 is presented in 142 Figure 3.8 The oxidation of l-(4-ethoxy-3-methoxyphenyl)propane (44) by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile 49 50 Figure 3.9 Proposed mechanism for alkyl-phenyl cleavage of phenolic model compounds 143 Figure 3.10 Possible mechanism for the formation of compound 48 Figure 3 . 1 0 . Veratryl alcohol is metabolized by P. chrysosporium via quinones and ring cleavage products.13 While the importance of Ca-hydroxylation to lignin biodegradation is not clear, it is quite possible that the demethoxylation and aromatic ring cleavage reactions found in this study are more important reactions than the Ca-hydroxylation previously reported for this model substructure.24,25 The phenylpropane substructures of lignin is probably also metabolized by P. chrysosporium through quinone and ring cleaved intermediates. 3.1.3. Oxidation of phenylpropene model compounds R 0CH3 v 51 R = CH 2OH or CHO Phenylpropene (51) is also a substructure of lignin22 and it has been reported that hydroxylation (diol formation) of the C^Cg double bond of l-(4-ethoxy-3-methoxyphenyl)-l-propene (52) by lignin peroxidase gives l-(4-ethoxy-3-methoxyphenyl)p^opane-l,2-diol24;^•28 (53). Diol formation from the C a - C 8 double bond was found to be the first step in the ctegradation of lignin model compounds by P. chrysosporium?9'32 1 4 5 52 53 54 Figure 3.11 The oxidation of l-(4-ethoxy-3-methoxyphenyl)-l-propene (52) by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile As shown in Figure 3.11, TDCSPPFeCl was able to oxidize 52 to give 4-ethoxy-3-methoxybenzaldehyde (24% yield), and 53 (33% yield) in the presence of t-BuOOH in aqueous acetonitrile. In addition, an acetonitrile incorporated product 54 (18% yield) was isolated. It has been previously found that the oxygen of the B-hydroxy group of 53 was derived from molecular oxygen,33 indicating a cation radical mechanism of the reaction. The formation of 54 certainly needs a CB-centred radical to be involved, supporting a cation radical mechanism of the reaction. Cytochromes P-450 are monooxygenases and function by activating molecular oxygen.34 They can catalyze diol formation from alkenes via an epoxide intermediate. Lignin peroxidases are peroxidases and have peroxidase and haloperoxidase activity.35 No monooxygenase activity of lignin peroxidase has been reported. Though the fact that P. chrysosporium can metabolize lindane36 (1,2,3,4,5,6-hexachlorocyclohexane) suggests the presence of other enzyme activities, it is still not clear which enzyme(s) is responsible for this reaction. Since lignin has a three-dimensional, heterogeneous structure, diol formation from lignin via an epoxide intermediate is unlikely. In order for oxygen atom to transfer and epoxide to be formed, the double bond of the substrate needs to approach closely the active site of the enzyme. It is unlikely that lignin as a substrate will meet this steric requirement. As both TDCSPPFeCl and alkene 52 are 146 small molecules, approach of the C a-CB double bond of 52 to TDCSPPFeCl is still possible, the formation of diol 53 from 52 via a epoxide intermediate cannot be excluded. The formation of 54, however, certainly suggests radical formation for at least one of the pathways of the oxidation of 52. i Proposed in Figure 3.12 is the mechanism for the formation of 53 and 54. One electron oxidation of 52 gives a cation radical, reaction of which with water gives a Cfl-centred radical. Reaction of the radical with dioxygen or further one electron oxidation followed by reacting with water gives the diol 53. Coupling of the Cfi-centred radical with an acetonitrile derived radical gives the acetonitrile incorporated product 54. When 52 was oxidized under highly aerobic conditions (bubbling oxygen using a sintered glass frit), litde of the solvent incorporated product 54 was detected, probably because of the competitive coupling of molecular oxygen with the CB-centred radical. 3.1.4. Oxidation of diarylpropane model compound Diarylpropane model compounds (B-l) are frequendy used substrates for lignin biodegradation studies. Oxidation of l-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)propane-l,3-diol (55) by lignin peroxidase gives mainly Ca-C f i cleavage products.25,28,33 Oxidation of 55 by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile (pH 3) gave 4-ethoxy-3-methoxybenzaldehyde (56, 40% yield), 4'-methoxy-a-hydroxyacetophenone (57, 12% yield), and 4-methoxybenzaldehyde (60, 20% yield) as major products (Figure 3.13). Small amounts of 4-ethoxy-3-methoxybenzoic acid (59) and 4-methoxybenzoic acid (61) were also isolated by preparative TLC and detected by GC and GCMS after methylation with diazomethane of the acidic products. 4-147 OEt Figure 3.12 Possible mechanism for the oxidation of l-(4-ethoxy-3-methoxyphenyl)propene 148 Figure 3.13 The oxidation of l-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)propane-l,3-diol (55) by TDCSPPFeCl and t-BuOOH Methoxyphenylglycol (58), which was a product in the oxidation of 55 by lignin peroxidase,25,28,33 was not isolated in this study. Careful G C and GCMS analyses of the acetylated products showed the presence of only trace amount of 4-methoxyphenylglycol diacetate. Oxidation of synthetic 58 under the condition used for the oxidation of 55 gave 57, 60, and a small amount of 61. Competitive oxidation of a 1:1 (molar ratio) mixture of 55 and 58 indicated that the B - l dimer 55 instead of the glycol 58 was preferentially oxidized under the conditions of this experiment The glycol 58 was therefore neither a major product nor an important intermediate for the oxidation of 55 by TDCSPPFeCl and t-BuOOH. The ketone 57 was formed directly. During the oxidation of 55 by lignin peroxidase,25,28'33 glycol 58 instead of the ketone 57 was the major product of the reaction while under anaerobic condition, the ketone 57 was not found. Oxidation of 55 by TDCSPPFeCl and t-BuOOH under anaerobic conditions also gave 58 as a major product and little ketone 57 was found. This suggests that the differences in the product distribution for the degradation of 55 by lignin peroxidase and by TDCSPPFeCl under aerobic condition was not the result of mechanistic differences. Experimental conditions were most probably responsible. It was noticed that when B- l model compounds were oxidized by TPPFeCl 3 7 or protohemin,38 the glycol, analogous to 58, was produced in smaller amounts than the corresponding ketone, suggesting that common differences exist between the reaction catalyzed by lignin peroxidase and those catalyzed by various biomimetic catalysts. 3.1.5. Oxidation of arylglycerol-B-aryl ether (B-O-4) model compounds The arylglycerol-B-aryl ether (B-O-4) substructure (4) is the most abundant substructure in lignin and in spruce lignin about 48% of the linkages connecting the 150 phenylpropanoid units are of the B-O-4 type. B-O-4 model compounds are one of the most frequendy used model compounds in lignin biodegradation studies. Oxidation of 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether (62) by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile (pH 3) gave 4-ethoxy-3-methoxybenzaldehyde (35% yield) and guaiacol (63, 30% yield) as the major products Figure 3.14 The oxidation of 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether by TDCSPPFeCl and t-BuOOH (Figure 3.14). Other products were detected in very small amounts and have not been identified. In the degradation of B-O-4 model compounds by lignin peroxidase,27,40 C a -C B cleavage was also the major reaction. 3.1.6. Oxidation of phenylcoumaran (B-5) model compounds The phenylcoumaran substructure (B-5, 8) constitutes about 9-12% of the substructures in spruce lignin.39 The degradation of B-5 model compounds by lignin peroxidase has not been reported. In order to investigate the ability of TDCSPPFeCl in degrading the various substructures of lignin, we studied the oxidation of an easily 151 available 6-5 dimer, 4-O-ethyldehydrodiisoeugenol (64) by TDCSPPFeCl and t-BuOOH. Oxidation of 64 by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile (pH 3) gave the side chain cleavage product 65 (-70% isolated yield) as the first product (Figure 3.15), further oxidation of which gave C a - C f i cleavage products 4-ethoxy-3-methoxybenzaldehyde and 70, acid 66, and aromatic ring cleavage products 67, 68, and 69. It was not clear if the ring opened acid 67 was a product of aromatic ring cleavage of acid 66 or was from the oxidation of the ring-cleaved aldehyde 68. The configuration of the double bonds in the ring cleaved products was elucidated by examining the coupling constants and by using NOE (nuclear overhauser effect) difference spectra as described in the experimental section. Degradation of B-5 model compounds by the ligninolytic culture of P. chrysosporium has been studied.41,42 In the oxidation of methyl dehydrodiconiferyl alcohol,41 the double bond of the side chain was first hydroxylated to give the glycol, which was further cleaved to give an aldehyde. In the oxidation of 64 by TDCSPPFeCl and t-BuOOH, the glycol was not isolated, probably because of its rapid oxidation. Aldehyde 65 was isolated as the first product. As described above, we showed by using l-(4-ethoxy-3-methoxyphenyl)-l-propene (52, Figure 3.11) that TDCSPPFeCl was able to catalyze the diol formation. Aromatic ring cleavage of veratryl alcohol and B-O-4 model compounds43-47 has been reported. It is interesting to note that in the case of B-O-4 dimers, all the ring cleavage products resulted from B-ring cleavage while cleavage of the A-ring has not been found. In addition, ring cleavage of B- l model compounds has never been reported. It is possible that the C a - C 6 cleavage reaction of B- l and B-O-4 model compounds occurs so quickly that the A-ring cation radical is not sufficiently long lived for aromatic ring cleavage to occur. The cation radicals of methoxybenzenes48 and of 152 Figure 3.15 The oxidation of 4-O-ethyldehydrodiisoeugenol (64) TDCSPPFeCl and t-BuOOH 153 the B-ring of a a-oxo B-O-4 model compound18 have been detected by ESR. Spectral observation of the A-ring cation radical of B- l and B-O-4 model compounds has not been reported. The isolation of A-ring cleavage products from 64 suggested that the C a - C B cleavage of this B-5 model compound is not the preferred route compared to that of B- l and B-O-4 model compounds. The B-ring of 65 is less readily oxidized than the A-ring due to the deactivation by the aldehyde group on the B-ring. 3.1.7. Oxidation of 5-5' biphenyl model compound The 5-5' biphenyl structure (6) is another main substructure of lignin and 9.5-11% of the linkages connecting the phenylpropanoid units of spruce Ugnin are of the biphenyl type.39 The guaiacyl moieties (71) of Ugnin are easily condensed by the 71 phenol oxidizing enzymes of white-rot fungi to give the 5-5' biphenyl structure,49,50 which is resistant to microbial attack.51 The abiUty of an enzyme or a biomimetic catalyst to degrade this substructure is therefore very important to Ugnin biodegradation. The metabolism of dehydrodivanillic acid (72) by bacteria52 and by P. chrysosporium5* has been studied and 5-carboxyvanillic acid 73 was found as a product in both cases (Scheme 3.2). The mechanism and enzyme(s) responsible for the formation of 73 is still not clear. The degradation of tetrameric model compounds 154 COOH COOH COOH COOH Scheme 3.2 containing the biphenyl structure by bacteria54,55 and by P. chrysosporium56 has also been studied. The major reactions were side chain cleavage, no aromatic ring cleavage or biphenyl bond cleavage was found. We selected 4,4'-diethyldehydrcdivanillin (74) as a biphenyl model compound to avoid complex side chain reactions. Oxidation of 74 by TDCSPPFeCl and t-BuOOH in aqueous acetonitrile (pH 3) gave acids 75 and 76 from side chain oxidation and aromatic ring cleavage products 77 and 78 from aromatic ring oxidation (Figure 3.16). It has been proposed57,58 that the aromatic aldehyde moiety of lignin cannot be metabolized direcdy by Ugnin degrading enzymes. It is reduced to an alcohol12,57 or oxidized to a acid57,58 before undergoing further metabolism. The fact that ring cleavage occurred only on the ring bearing a carboxyl group certainly supports the second pathway of aromatic aldehyde metabolism. Direct aromatic ring cleavage products similar to 77 and 78 have been found in this study and reported in the literature17. The similarities of these reactions suggest that they follow the same mechanism, that is, initiation by an one-electron oxidation process. The degradation of biphenyl model compounds by lignin peroxidase has not been reported. The formation of aromatic ring cleavage products 77 and 78, probably 155 ON CHO CHO CH3O " ^ r " ^ Y ^ O C H 5 OEt OEt 74 COOHCHO CHO COOH CH3O + OCH- CH3O OEt C 0 2 C H 3 C 0 2 E t COOH COOH CH3O + 77 COOH COOH OCH- CH3O OEt 78 C 0 2 C H 3 C 0 2 E t Figure 3.16 The oxidation of 4,4'-diethyldehydrodivanillin (74) by TDCSPPFeCl and t-BuOOH through an one-electron oxidation pathway, suggests that lignin peroxidase might also oxidize the biphenyl substructure of Ugnin. Veratryl alcohol is metaboUzed to carbon dioxide by P. chrysosporium via quinone and ring cleavage product intermediates.12 No demethoxylation products of the biphenyl model compound 74 were found in this study. Aromatic ring cleavage is probably the first and most critical step for the biodegradation of biphenyl substructures of Ugnin. 3.1.8. Oxidation of pyrene In addition to a wide variety of lignin model compounds and polychlorinated phenols, lignin peroxidase can also catalyze the oxidation of polycycUc aromatics. Pyrene (79) was oxidized to give pyrene-1,6-dione (80) and pyrene-1,8-dione (81) through a one-electron oxidation mechanism.59 Benzofajpyrene59,60 was also oxidized by lignin peroxidase to give benzo[a]pyrene 1,6-, 3,6-, and 6,12- quinones. The ability of TDCSPPFeCl to catalyze the oxidation of polycycUc aromatics was studied using pyrene as a substrate. Pyrene was oxidized rapidly by TDCSPPFeCl/mCPBA at pH 3 to give pyrene-1,6-dione and pyrene-1,8-dione as major products (Figure 3.17). Further oxidation of 0 o 0 80 81 Figure 3.17 The oxidation of pyrene by TDCSPPFeCl and mCPBA 157 the quinones by TDCSPPFeCl/mCPBA was not successful as the quinones were not readily oxidized and TDCSPPFeCl was not stable towards mCPBA at pH 3. The oxidation of pyrene-l,8-dione by /nCPBA and Cli 6 TSPPMnCl, which is a more stable catalyst than TDCSPPFeCl, gave mainly black tars which was insoluble in either water or organic solvents. 3.1.9 Oxidation of 2,4,6-trichIorophenol It has been reported61 that lignin peroxidase could also catalyze the oxidation of chlorinated phenols. Pentachlorophenol and 2,4,6-trichlorophenol was oxidized61 to give tetrachloro-p-benzoquinone and 2,6-dichloro-p-benzoquinone, respectively. We have shown that TDCSPPFeCl is able to oxidize 2,4,6-trichlorophenol (82) to give 2,6-dichloro-p-benzoquinone (83) as a major product along with an unidentified product (Figure 3.18). 3.2 Conclusion It has been shown that TDCSPPFeCl as a biomimetic catalyst can mimic the function of lignin peroxidase in the oxidation of a variety of lignin model compounds C l 0 82 83 Figure 3.18 The oxidation of 2,4,6-trichlorophenol by TDCSPPFeCl and mCPBA. 158 OH 0 and related substrates, mcluding veratryl alcohol, veratryl acetate, phenylpropane and phenylpropene model compounds (44 and 52), B-l (55), B-O-4 (62), B-5 (64), and 5,5'-biphenyl (74) model compounds. Pyrene and 2,4,6-trichlorophenol can also be oxidized by the iron porphyrin catalyst, suggesting that other environmental pollutants might also be degraded by this catalyst The isolation of direct ring cleavage products from the oxidation of veratryl acetate, l-(4-ethoxy-3-methoxyphenyl)propane (44), 4-O-ethyldehydVodiisoeugenol (64), and 4,4'-dlethyldehycU,om\anillin (74) supports the mechanism for aromatic ring cleavage of lignin model compounds as shown in Figure 3.3. The fact that methanol was incorporated only into the Q-position of veratryl alcohol to give methanol incorporated products 37 and 38 provides evidence that the cation radical of veratryl alcohol is preferentially attacked by solvent at its exposition. As shown in Figure 3.3, attack of water onto the cation radical is the first step for both the aromatic ring cleavage and the demethoxylation reactions. The formation of the acetonitrile incorporated product 54 in the oxidation of the phenylpropene model compound 52 requires the involvement of a carbon centred radical, suggesting a cation radical mechanism for the oxidation of this type of substructure. The oxidation of the phenylcoumaran (B-5) model compound 64 to give C a-C 6 cleavage and aromatic ring cleavage products suggests that the B-5 substructure can be degraded in a manner similar to other substructures of Ugnin, probably through the same cation radical mechanism. For the first time aromatic ring cleavage products derived from the A-ring of a dimeric model compound have been isolated. A 5,5'-biphenyl model compound (74) which represents the Ugnin substructure most resistant to microbial degradation can be oxidized by TDCSPPFeCl, suggesting 159 that the iron porphyrin is powerful enough to oxidize all the major substructures of lignin. The oxidation of 5,5'-biphenyl model compounds by purified lignin peroxidase has not been reported. The formation of aromatic carboxylic acids from TDCSPPFeCl catalyzed oxidation of the biphenyl model compound 74 and the B-l model compound 55 suggests that the iron porphyrin is probably a more powerful catalyst than lignin peroxidase. 33 Experimental 3.3.1 General Equipment. *H NMR spectra were recorded on Varian XL 300 or Bruker WH 400 spectrometers using tetramethylsilane as internal standard. 13C NMR spectra were recorded on Varian XL 300 spectrometer. Low resolution and high resolution mass spectra were recorded on an AE1 MS9 and a Cratos MS 50 spectrometer, respectively. A HP 5890A Gas Chromatograph equipped with a 25 meter HP 17 capillary column was used for all the GC analyses. A Delsi Nermag RIO-lOC/Kratos MS80RFA combination was used for the Gas Chromatography-Mass Spectroscopy (GCMS) analyses. Infrared spectra were obtained on a Perkin-Elmer 1710 Fourier Transform spectrometer. Preparative thin layer chromatography (TLC) were carried out on a Model 7924T Chromatotron (Harrison Research) using silica gel PF^ as absorbent. The melting points (uncorrrected) were deterrnined on a Bristoline 6548-J17 Microscope equipped with a Thomas Model 40 Micro Hot Stage. Materials. Merck silica gel 60 (70-230 mesh) was used for column chromatography. Merck precoated silica gel PF^ TLC plate (0.2 mm) was used for 160 analytical T L C analyses. Veratryl alcohol, eugenol, isoeugenol, t-BuOOH, 4'-methoxyacetophenone, and pyrene were from Aldrich. Vanillin and mCPBA was purchased from Fisher Scientific Company and 2,4,6-trichlorophenol was from Eastman Organic Chemicals. TDCSPPFeCl and Cl 1 6 TSPPMnCl were synthesized as described in Chapter 2. 3.3.2 Syntheses of lignin model compounds Veratryl acetate was prepared by acetylation of veratryl alcohol (0.84 grams, 5 mmole) with excess 1:1 acetic anhydride/pyridine at room temperature for 24 hours. Saturated sodium bicarbonate (5 mL) was then added to the mixture and stirred for 20 minutes, extracted (2x15 mL) with ethyl acetate. The ethyl acetate was combined, washed in sequence with saturated sodium bicarbonate (5 mL), water (5 mL), and 10% HC1 (2x5 mL). The ethyl acetate layer was dried over sodium sulphate, evaporated, and purified by column chromatography (silica gel, 7:3 hexanes/ethyl acetate as eluent). The veratryl acetate obtained was characterized by ' H N M R and MS. 4-Ethoxy-3-methoxybenzyl alcohol was prepared as reported.29 l-(4-ethoxy-3-methoxyphenyl)propane (44) was synthesized by hydrogenation of eugenol in ethanol by hydrogen/Pd (on charcoal, 1%) followed by ethylation with ethyl iodide and sodium hydride in N,N-dimethylformamide (DMF). Eugenol (3.28 grams, 0.01 moles) and Pd (1% on charcoal, 0.03 g) were added to ethanol (20 mL) and stirred at room temperature for overnight under hydrogen (1 atm.). The mixture was filtered, and evaporated to give 2-methoxy-4-propylphenol in quantitative yield. The crude 2-methoxy-4-propylphenol was subjected to ethylation without purification. Sodium hydride (2.8 g, 50%) and ethyl iodide (4 g, 0.026 moles) were added to D M F 161 (10 mL) containing 3 g of 2-methoxy-4-propylphenol and stirred for 60 minutes. Water (10 mL) was added to the solution to decompose remaining sodium hydride and the solution was then extracted twice with ethyl acetate (20 mL). The ethyl acetate was evaporated and the residue purified by column chromatography using 80:20 petroleum ether (high boiling):ethyl acetate as eluent. l-(4-Ethoxy-3-methoxyphenyl)propane (44) was obtained as colourless flakes in 85% yield after recrystallization from diethyl ether/hexane. l-(4-Ethoxy-3-methoxyphenyl)-l-propene (52) was synthesized by ethylation of isoeugenol with ethyl iodide and sodium hydride in D M F following the same procedures as described above. The product was obtained in 80% yield. 4-Ethoxy-3-methoxyphenylglycerol-fi-guaiacyl ether62 (62), l-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)propane-l,3-diol6i (55), ethyl dehydrodiisoeugenol6* (64), and 4,4'-diethyldehydrodivanillin65 (74) were synthesized following reported procedures. 4-Methoxyphenylglycol (58) was synthesized following a reported procedure66. 4'-Methoxyacetophenone (3.0 g) was reacted with cupric bromide (80 mesh, 9.0 g) in 1:1 ethyl acetate/chloroform mixture (40 mL) at reflux temperature for 3 hours to give 4'-methoxy-a-bromoacetophenone in 90% yield. The product was purified by column chromatography using 1:99 (volume) methanokmethylene chloride as eluent to give a light yellow crystaline product, reaction of which with sodium acetate (3.7 g) in acetic anhydride (10 mL) at reflux temperature for 2 hours gave 4'-methoxy-oc-hydroxyacetophenone acetate in 80% yield. Reduction of the resulting 4'-methoxy-oc-hydroxyacetophenone acetate by sodium borohydride (2.0 g) in methanol (20 mL) at 0°C for 3 hours gave 4-methoxyphenylglycol (58) in 90% yield. 162 33.3 The oxidation of the lignin model and related compounds The TLC separation of reaction mixtures. The products were separated on a Chromatotron using either methanol/methylene chloride (0-1% methanol, volume) or ethyl acetate/hexane (10-30% ethyl acetate, volume) as eluent. The purity of each components was checked by GC and analytical TLC. Impure components were further separated on the Chromatotron. The yields of the products were based on substrate and estimated from GC integration area. All the reactions were carried out in air at room temperature unless otherwise indicated. No products were detected by TLC in control experiments without the iron porphyrin catalyst. The oxidation of veratryl alcohol and veratryl acetate. The substrate (0.3 mmole) and TDCSPPFeCl (0.5 pjnole in 0.5 mL water) were dissolved in either pH 3 phosphate buffer (0.1 M) or methanol (20 mL) and mCPBA (0.6 mmole) was added as a powder and the solution was stirred at room temperature for 2 hours. The solution was partitioned between saturated sodium bicarbonate (20 mL) and ethyl acetate (20 mL). The aqueous layer was further extracted twice with ethyl acetate (10 mL) and the organic extracts were combined, dried over anhydrous sodium sulphate, and evaporated. When veratryl acetate was the substrate, the products were analyzed by GC and separated on a Chromatotron. When veratryl alcohol was the substrate, the products could be separated on a Chromatotron with or without acetylation with 1:1 acetic anhydnd^ /pyridine. The sample for GC analysis had to be acetylated since veratryl alcohol was found to be oxidized on the GC column. The oxidation of veratryl alcohol in ethanol was carried out under similar conditions as that for the oxidation of veratryl alcohol in methanol except a mixture of 5 mL DMF (to increase the solubility of TDCSPPFeCl) and 20 mL ethanol was used as 163 the solvent. The condition for the oxidation of 4-ethoxy-3-methoxybenzyl alcohol was the same as that for veratryl alcohol. The hydrolysis of 2-hydroxymethyl-4,4\5-trimethoxy-2,5-cyclohexadiene-l-one (37). 0.25 grams of 37 was dissolved in 10 mL of 1:1 THF/water, shaken with 2 mL concentrated H C l , and extracted with ethyl acetate. The ethyl acetate was washed with water (5 mL), dried and evaporated. The product was shown to be pure 30 as characterized by *H N M R and MS. The oxidation of l-(4-ethoxy-3-methoxyphenyl)propane (44). 0.25 mmole of 44 was oxidized by t-BuOOH (1.25 mmole) and TDCSPPFeCl (0.5 umole) in 20 mL of 1:1 acetonitrile/pH 3 phosphate buffer at room temperature in air for 3 hours. Unreacted t-BuOOH was decomposed by adding solid potassium iodide. The solution was then extracted with ethyl acetate (3x10 mL) and the iodine in the extract was removed by washing with aqueous sodium thiosulphate. The products were acetylated with 1:1 acetic anhydride/pyridine before being subjected to GC, GCMS analyses and separation on a Chromatotron. The yield (estimated from GC) of the products 45, 46, 47, and 48 was 15%, 23%, 20%, and 20%, respectively. The oxidation of 1 -(4-ethoxy-3-methoxyphenyl)-l-propene (52) was carried out under the same conditions as that for 44 except the reaction time was 2 hours. The oxidation of 4-ethoxy-3-methoxyphenylglycerol-fi-guaiacyl ether (62). 0.5 mmole of 62 was oxidized by 1 umole of TDCSPPFeCl and either 1.5 mmole of mCPBA or 2.5 mmole of t-BuOOH in 50 mL of 1:1 acetonitrile/pH 3 buffer for 3 hours at room temperature. The workup and analysis procedure was the same as that for 44. The conditions for the oxidation of l-(4-ethoxy-3-methoxyphenyl)-2-(4-164 methoxyphenyl)propane-l £-diol (55) were the same as those for 62 except the reaction time was 6 hours. The products were separated into neutral and acidic fractions by extracting the ethyl acetate layer with saturated sodium bicarbonate. The acidic part was methylated with diazomethane which was generated by using p-toluenesulfonylmethylnitrosamide as reported.67 The neutral part was acetylated and both the methylated products and the acetylated products were analyzed as described above. The oxidation of 55 under anaerobic condition. A 25 mL solution (1:1 acetonitrile/pH 3 buffer) containing 55 (0.25 mmole) and TDCSPPFeCl (0.5 pimole) in a 50 mL Schlenk tube was degassed by means of freeze-thaw (3 cycles). The oxidant (mCPBA, 0.75 mmole) was put into another small Schlenk tube (10 mL) connected with the one containing the solution and similarly degassed. The contents of the two Schlenk tubes were mixed and stirred for 6 hours after being warmed up to room temperature. The products were acetylated by excess 1:1 acetic anhydride/pyridine and analyzed by G C and G C M S . The ^ oxidation of ethyldehydrodiisoeugenol (64) (0.5 mmole) by TDCSPPFeCl (0.5 umole) and t-BuOOH (2.5 mmole) in 2:1 acetonitrile/pH 3 buffer (60 mL) for 2 hours in air gave 65 as the major product along with small amount of 4-ethoxy-3-methoxybenzaldehyde. Compound 65 was separated using a Chromatotron and further oxidized under the same conditions for 3 hours. The oxidation of 65 was found to give both neutral and acidic products but the separation of the two groups of compounds by sodium bicarbonate extraction was not successful due to the large molecular sizes of the products. The products were thus methylated with diazomethane before separation. The products were very difficult to separate on a Chromatotron and each component from first separation was further purified on the Chromatotron. Two 165 parallel experiments were carried out to obtain enough material for N M R and MS characterization of the products. The oxidation of 4,4''-diethyidehydrodivaniliin (74). 0.5 mmole of 74 was oxidized by TDCSPPFeCl (1 umole) and t-BuOOH (0.25 mmole) in 1:1 acetonitrile/pH 3 buffer (60 mL) for 8 hours. The products were separated into neutral and acidic parts. The neutral part (- 60% of the starting material) contained only the starting material and the acidic part was methylated with diazomethane before separation on the Chromatotron. The oxidation of pyrene (0.5 mmole) by TDCSPPFeCl (0.1 umole) and mCPBA was carried out in 3:1 acetonitrile/pH 3 buffer (40 mL) at room temperature. The reaction completed in 30 minutes, as followed by UV-vis spectroscopy. Saturated sodium bicarbonate (30 mL) was added to the reaction mixture followed by extracting with ethyl acetate (3 x 50 mL). The products were separated by column chromatography (a large column, 4x30 cm, had to be used due to the low solubility of the compounds in the solvent) using 80:20 toluene/ethyl acetate as eluent. The further oxidation of one of the oxidation products of pyrene, pyrene-1,8-dione (10 mg), by Cl 1 6 TSPPMnCl (0.2 umole) and mCPBA (0.2 g) was carried out for 12 hours in 2:1 acetonitrile/pH 3 buffer (30 mL). The reaction mixture was partitioned between saturated sodium bicarbonate (20 mL) and ethyl acetate (20 mL). Black tars which were insoluble in either water or organic solvents were found. The small amounts of compounds in the ethyl acetate extract was not identified. The oxidation of 2,4,6-trichlorophenol. 2,4,6-Trichlorophenol (2 mmole) was oxidized by TDCSPPFeCl (0.5 umole) and mCPBA (4 mmole) in 2:1 acetonitrile/pH 3 buffer (30 mL) for 12 hours. The reaction mixture was extracted three times with ethyl acetate (3 x 30 mL). T L C analysis using methanol/methylene chloride (0.5:99.5) as 166 eluent showed the presence of only small amount of starting material. The products were separated on silica gel column using methanol/methylene chloride (0.5:99.5) as eluent to give a mixture of two yellow compounds, which was further separated on column using ethyl acetate/hexane (5:95) as solvent. 3.3.4 Characterization of compounds A l l compounds were characterized by ' N M R and MS unless otherwise specified. Elemental analyses and 1 3C N M R were performed when enough product was available. Methyl (z)-furan-5(2H)-one-3-butenoate (26) OH N M R and MS the same as those reported11) ' H N M R (CDC13) 8: 3.78 (3H, S), 5.25 (2H, m), 6.13 (IH, d, J=12Hz), 6.30 (IH, m), 6.77 (IH, dd, J,=12Hz, J2=l-2 Hz). MS m/z(%): 168 (12.6, M), 137 (19.4), 136(48.2), 124(35.2), 111(26.0), 109(33.4), 108(21.6), 81(49.2), 80(28.6), 79(60.5), 59(22.5), 53(94.8), 52(45.5), 51(100.0), 50(46.6). Methyl (Z)-6-oxo-2H-pyran-3(6H)-vlideneacetate (28) ( ! NMR and MS similar to those reported17) ' H N M R (CDCI3) 5: 3.80(3H, S), 5.03(2H, d, J=2Hz), 5.94QH, m), 6.21(1H, dd, J,=10Hz, J2=2Hz), 8.35(1H, dd, J ^ l H z , J2=10Hz). MS m/z(%): 168(13.0), 136(17.9), 124(14.8), 112(10.7), 109(33.0), 108(14.0), 167 81(36.5), 80(10.7), 79(10.9), 59(12.6), 53(100.0), 52(23.3), 51(37.0), 50(19.0) M.P.: 87-90°C (reported17: 87-90°C). Methyl (E)-6-oxo-2H-pvran-3(6H)-vlideneacetate (29)(1H N M R and MS similar to those reported17) ! H N M R (CDC13) 8: 3.78(S, 3H), 5.69(2H, d, J=3Hz), 5.99(1H, m) 6.19(1H, d, J=10Hz) 7.01(1H, dd, J ^ l H z , J2=10Hz). MS m/z(%): 168(14.0), 137(16.7) 136(100), 111(13.5), 109(25.2), 108(37.3), 97(11.2), 83(10.5), 81(42.3), 80(31.8), 79(29.1), 53(61.0), 52(20.1), 51(37.2). M.P.: 147-150°C (reported17: 147-150 °C). 2-Hvdroxvmethvl-5-methoxv-2,5-cvclohexadiene-l,4-dione (30)(1H N M R and MS similar to those reported68) •H N M R (CDCI3) 8: 3.85 (3H, S), 4.57(2H, J=l Hz), 5.92(1H, S), 6.72(1H, t, J=l Hz). 1 3 C N M R (CD3OD) 8: 188.798, 183.500, 160.461, 150.891, 129.456, 108.426, 58.959, 56.959. MS m/z(%): 170(M, 1.2), 169(16.5), 168(100.0), 167(64.9), 153(16.9), 150(12.2), 168 140(10.2), 139(55.4), 122(15.6), 111(31.9), 83(14.4). Exact mass calcd. for C,HiA : 170.0579; found (HRMS): 170.0588. M.P.: 142-143°C (reported68: 144-145°C). 4,5-Dimethoxy-3,5-cyclohexadiene-1,2-dione (32)C'H N M R and MS similar to those reported17) *H N M R (CDC13) 8: 3.90(6H, S), 5.78(2H, S). MS m/z(%): 170(5.74), 168(3.38), 167(7.28), 155(4.12), 149(24.44), 140(100.0), 125(27.94), 69(18.34). Exact mass calcd. for QFLCv 168.0420; found (HRMS): 168.0403. 2-Acetoxvmethyl-5-methoxv-2,5-cyclohexadiene-1,4-dione (34) ' H N M R (CDCI3) 8: 2.17(3H, S), 3.85(3H, S), 5.02(2H, d, J=l-2Hz), 5.95(1H, S), 6.62(1H, t, J=l-2Hz). 1 3C N M R (CDCI3) 8: 185.947, 181.402, 169.860, 158.628, 143.815, 129.264, 107.495, 59.642, 56.468, 20.774. MS m/z(%): 210(2.41), 168(100.0), 167(25.23), 153(10.16), 139(13.23), 69(22.9), 169 43(78.0). Exact mass calcd. for C10fil0Os: 210.0528, found (HRMS): 210.0535 M.P.: 158-160°C. 3,4-Dimethoxyphenol (35) ' H N M R (CDC13) 8: 3.84(3H, S), 3.86(3H, S), 4.56(1H, broad), 6.36(1H, dd, J,=9Hz, J2=2Hz), 6.50(1H, d, J=2Hz), 6.75(1H, d, J=9Hz) MS m/z(%): 154(100.0), 139(94.8), 111(62.7), 96(10.6), 93(40.8), 81(11.0), 69(21.5), 68(11.1), 65(25.4), 55(12.7). Dimethyl 3-acetoxvmethyl-2,4-hexadienedioate (36) ' H N M R (CDCI3) 5: 2.12(3H, S), 3.705(3H, S), 3.710(3H, S), 4.85(2H, m), 6.00(1H, d, J=12Hz), 6.00(1H, d, J=l-2Hz), 7.06(1H, dd, J^UHz, J2=l-2Hz). MS m/z(%): 242(0.1), 184(9.8), 183(100.0), 169(28.4), 141(22.7), 136(10.5), 109(43.8), 43(55.5). Exact mass calcd. for C n H M 0 6 : 242.0786; found (HRMS): 242.0782. 2-Hvdroxvmethvl-4,4,5-trimethoxv-2,5-cvclohexadiene-1 -one (37) (identified as its acetate) 170 ' H N M R (CDCI3) 5: 2.14(3H, S), 3.34(6H, S), 3.84(3H, S), 4.92(2H, d, J=l-2Hz), 5.67(1H, S), 6.53(1H, t, J=l-2Hz). 1 3 C N M R (CDCI3) 8: 184.449, 170.076, 169.097, 136.119, 136.057, 103.832, 94.349, 60.314, 56.252, 51.534, 20.898. MS m/z(%): 256(2.2), 241(25.9), 199(33.2), 183(100.0), 182(25.9), 167(29.0), . 155(50.8), 139(29.7), 124(23.4), 123(61.5), 95(28.4), 69(32.9), 43(85.9). Anal, calcd. for C 1 2 H 1 6 0 6 , C: 56.25; H : 6.29. found: C: 56.37; H : 6.37. Exact mass calcd. for C 1 2 H 1 6 0 6 : 256.0948, found (HRMS) 256.0948. M.P.: 79-81°C. 2-Dimethoxvmethvl-4,4,5-trimethoxy-2,5-cvclohexadiene-1 -one (38) >H N M R (CDCI3) 8: 3.35(6H, S), 3.39(6H, S), 3.82(3H, S), 5.34(1H, S), 5.65(1H, S), 6.81(1H, S). 1 3 C N M R (CDCI3) 8: 184.744, 168.959, 137.144, 104.092, 97.798, 94.454, 56.222, 54.032, 51.540, 29.679. MS (EI) m/z(%): 243(0.3), 227(100.0), 212(19.8), 180(37.3), 153(10.8), 75(31.5), 69(12.1). 171 MS (DCI+NH3) m/z(%): 259(29.6, M+l), 227(58.4), 213(13.6), 212(100.0). 4-Ethoxy-2-hvcumvmethvl-4,5-fl (40) (isolated as its acetate) ' H N M R (CDC13) 8: 1.23(3H, t, J=7Hz), 2.13(3H, S), 3.31(3H, S), 3.40-3.60(2H, m, when decoupled at 1.23 , pattern close to a A B quartet), 4.91(2H, d, J=lHz), 5.66(1H, S), 6.52(1H, t, J=lHz). MS m/z(%): 270(1.5), 241(29.7), 199(35.8), 197(87.2), 183(100.0), 182(26.1), 169(45.2), 167(43.8), 155(32.4), 153(26.7), 141(31.0), 139(25.6), 123(51.3), 109(57.4), 95(24.2), 69(28.6), 43(88.5). 5-Ethoxv-2-hvdroxvmethvl-4,4-dirnethoxv-2,5-cvclohexadiene-1 -one (41) (identified as its acetate) ' H N M R (CDCI3) 8: 1.47(3H, t, J=8Hz), 2.13(3H, S), 3.33(6H, S), 4.05(2H, q, J=8Hz), 4.92(2H, d, J=lHz), 5.63(1H, S), 6.51(1H, t, J=lHz). MS m/z(%): 270(1.8), 199(23.8), 197(100.0), 182(32.9), 169(37.2), 153(24.6), 141(35.7), 139(34.3), 123(14.8), 109(43.9), 69(27.1), 43(76.0). 2-Dimethoxvrnethvl-5-ethoxv-3,4-dimethoxv-2,5-cyclohexadiene-1 -one (42) ' H N M R (CDC13) 8: 1.46(3H, t, J=8Hz), 3.35(6H, S), 3.39(6H, S), 4.03(2H, q, 172 J=8Hz), 5.34(1H, S), 5.61(1H, S), 6.79(1H, S). MS m/z(%): 242(13.3), 241(100.0), 226(28.4), 194(44.9), 181(17.4), 167(10.8), 153(16.2), 75(24.1), 69(13.6). 1 -(4-Ethoxv-3-methoxyphenvl)propane (44) lH N M R (CDC13) 8: 0.94(3H, t, J=8Hz), 1.44(3H, t, J=7Hz), 1.57-1.67(2H, m), 2.53(2H, t, J=9Hz), 3.86(3H, s), 4.07(3H, q, J=7Hz), 6.67-6.81(3H, m). MS m/z(%): 194(35.2), 166(10.9), 165(35.2), 138(10.6), 137(100.0), 122(6.5). M.P.: 33-34°C. (6)-Ethyl (l)-methyl 3-propyl-2,4-hexadienedioate (45) J H N M R (CDCI3) 8: 0.93(3H, t, J=7Hz), 1.26(3H, t, J=7Hz), 1.44-1.57(2H, m), 2.39(2H, t, J=7Hz), 3.67(3H, s), 4.34(2H, q, J=7Hz), 5.76(1H, s, broad), 5.88(1H, d, J=12Hz), 7.05QH, dd, J ^ H H z , J 2=l-2Hz). MS m/z(%): 226(2.74), 211(1.10), 195(4.73), 194(2.27), 181(9.78), 167(71.67), 153(100.0), 149(10.12), 139(75.41). Exact mass calcd. for C 1 2 H 1 8 0 4 : 226.1200; found (HRMS): 226.1204. 173 2-Ethoxv-5-propyl-2,5-cvclohexadiene-1,4-dione (46) l H N M R (CDCI3) 8: 0.97(3H, t, J=8Hz), 1.47(3H, t, J=7Hz), 1.51-1.57(2H, m), 2.41(2H, m), 4.01(2H, q, J= 7Hz), 5.90(1H, s), 6.49(1H, s, broad). MS m/z(%): 194(4.0), 179(3.7), 167(4.1), 166(5.1), 165(8.4), 151(34.8), 150(23.7), 149(32.6), 69(100.0), 67(22.3), 57(25.6), 55(23.4), 53(28.0), 44(20.6), 43(34.4), 41(30.0). l-(4-Ethoxv-3-methoxvphenyl)propanol (47) (identified as its acetate, its G C retention time was the same as that of the synthesized standard and its MS was similar to that of the standard. MS m/z(%) 252(30.1), 193(22.8), 192(28.0), 181(100.0), 165(15.5), 164(27.4), 163(14.7), 149(14.9), 133(12.1), 125(15.4), 107(14.9), 105(12.4), 103(10.9), 93(17.5), 91(15.7), 77(15.5), 65(13.1), 57(18.7), 43(54.3). 4-Hvdroxv-2-methoxv-4-propyl-2,5-cvclohexadiene- 1-one (48) ' H N M R (CDC13) 8: 0.94(3H, t, J=8Hz), 1.26-1.36(2H, m), 1.76-1.81(2H, m), 1.92(1H, s, exchangeable), 3.69(3H, s), 5.72(1H, d, J=2-3Hz), 6.20(1H, d, J=10Hz), 6.83(1H, dd, J1=2-3Hz, J2=10Hz). MS m/z(%): 182(13.9), 166(21.8), 140(22.8), 139(100.0), 111(39.3), 94(29.41), 93(33.3), 81(26.7), 79(32.0), 77(46.0), 69(88.0), 68(29.3), 67(25.5), 66(28.9), 65(68.4), 174 55(85.5), 53(53.4), 52(26.6), 51(47.6). Exact mass calcd. for C 1 0 H 1 4 O 3 : 182.0939; found (HRMS): 182.0942. FTIR (KBr pellet) (cm 1): 3436, 1672, 1645, 1214. 1 -(4-Ethoxv-3-methoxvphenyl)-1 -propene (52) [ H N M R (CDC13) 8: 1.45(3H, t, J=7Hz), 1.86(3H, dd, Jj=7Hz, J,=l-2Hz), 3.88(3H, s), 4.09(2H, q, J=7Hz), 6.06-6.15(lH, m), 6.32-6.37(lH, m), 6.79-6.91(3H, m). MS M/z(%): 192(100.0, M), 165(6.8), 164(63.1), 163(35.8), 149(31.0), 133(14.7), 131(17.2), 121(11.5), 107(34.6), 103(30.0). M.P.: 58-59°C (reported69: 64°C). l-(4-Ethoxv-3-methoxvphenyl)propane-l,2-diol (53) (isolated as its diacetate) ' H N M R (CDCI3) 8:1.08(3H, d, J=7Hz), 1.45(3H, t, J=7Hz), 2.05(3H, s), 2.07(3H, s), 3.89(3H, s), 4.10(2H, q, J=7Hz), 5.23-5.29(lH, m), 5.71(1H, d, J=8Hz), 6.82-6.91 (3H, m). MS m/z(%): 310(8.0), 266(3.0), 250(9.0), 223(7.5), 208(15.4), 182(11.3), 181(100.0), 151(19.6), 125(10.1), 43(41.6). 175 Exact mass calcd. for QgH^Cv 310.1414; found (HRMS): 310.1416. 4-Hvdroxv-4-(4-ethoxy-3-memoxvr)henvl)-3-memvlbutvronitrile (54) (identified as its acetate) ' H N M R (CDC13) 8: 1.00(3H, d, J=7Hz), 1.45(3H, t, J=8Hz), 2.10(3H, s), 2.30-2.40(2H, m), 2.46-2.53(lH, m), 3.89(3H, s), 4.10(2H, q, J=8Hz), 5.51 and 5.60 (IH, two doublets in a ratio of 7:2, J=8Hz), 6.80-6.84(3H, m). MS m/z(%): 291(10.1), 232(3.4), 223(3.6), 204(1.4), 203(1.8), 182(10.6), 181(100.0), 164(3.7), 153(14.5), 86(17.3), 84(27.4), 43(35.3). Exact mass calcd. for C 1 6 H 2 A N : 291.1465; found (HRMS): 291.1471. l-(4-Ethoxv-3-methoxvphenvl)-2-(4-methoxvphenvl)propane-l,3-diol (55) (characterized as its diacetate, mixture of stereo isomers). ' H N M R (CDCI3) 8: 1.38-1.47(3H, two triplets centred at 1.40 and 1.44, respectively), 1.94-2.10(6H, four singlets at 1.94, 1.96, 2.01, and 2.09, respectively), 3.32-3.45(lH, m), 3.70-3.79(6H, four singlets at 3.71, 3.75, 3.76, and 3.79, respectively), 4.00-4.13(2H, m), 4.25-4.51(2H, m), 5.94-6.05(lH, two doublets centred at 5.96 and 6.04, respectively), 6.49-7.11(7H, m). MS m/z(%): 416(2.9, M), 356(2.3), 297(1.9), 223(23.2), 182(11.1), 181(100.0), 134(12.7), 43(23.0). 176 4-Ethoxv-3-methoxybenzaldehvde (56) ' H N M R (CDCI3) 8: 1.51(3H, t, J=7Hz), 3.94(3H, s), 4.20(2H, q, J=7Hz), 6.98(1H, d, J=8Hz), 7.42-7.48(2H, m), 9.86(1H, s). MS m/z(%): 181(6.6, M+l), 180(62.0, M), 152(55.1), 151(100.0), 109(11.2). M.P.: 59-60°C (reported70: 64-65°C) 4/-Methoxy-a-hvdroxvacetophenone (57) ] H N M R (CDCI3) 8: 3.58(1H, t, J=5Hz, exchangeable), 3.90(3H, s), 4.83(2H, d, J=5Hz), 6.98(2H, d, J=9Hz), 7.90(2H, d, J=9Hz). MS m/z(%): 166(10.0), 136(30.7), 135(100.0), 107(35.5), 92(46.2), 77(64.4), 64(21.0). M.P.: 101-103°C (reported71: 100°C, 104°C). A'-Methoxyphenyl glycol (58) ! H N M R (CDC13) 8: 2.17(1H, broad, exchangeable), 2.55(1H, broad, exchangeable), 3.60-3.79(2H, m), 3.81(3H, s), 4.78(1H, m), 6.90(2H, d, J=8Hz), 7.31(2H, d, J=8Hz). 177 MS m/z(%): 168(1.1, M) , 150(14.5), 137(14.4), 136(14.9), 135(29.5), 122(10.6), 121(100.0), 91(13.7), 78(20.7), 77(43.7). M.P.: 79-80°C (reported72: 82°C) 4-Ethoxy-3-methoxvbenzoic acid (59) (isolated as its methyl ester) *H N M R (CDC13) 6: 1.52(3H, t, J=7Hz), 3.90(3H, s), 3.93(3H, s), 4.16(2H, q, J=7Hz), 6.88(1H, d, J=8Hz), 7.54(1H, d, J=2Hz), 7.66(1H, dd, J^SHz, J2=2Hz). MS m/z(%): 211(5.8, M+l), 210(4.2, M) , 182(40.4), 179(10.6), 152(11.3), 151(100.0), 149(19.3), 123(15.0), 79(19.6), 77(11.1), 65(14.0). 4-Methoxybenzaldehvde (60) ' H N M R (CDCI3) 8: 3.93(3H, s), 7.05(2H, d, J=8Hz), 7.88(2H, d, J=8Hz), 9.62(1H, s). MS m/z(%): 137(6.1, M+l), 136(73.2, M), 107(15.0), 92(12.7), 77(28.3). 4-Methoxybenzoic acid (61) (isolated as its methyl ester) ' H N M R (CDCI3) 5: 3.88(3H, s), 3.90(3H, s), 6.93(2H, d, J=8Hz), 8.00(2H, d, J=8Hz). 178 MS m/z(%): 167(3.5, M+l), 166(8.9, M), 135(46.0), 107(10.1), 85(27.7), 83(40.2), 77(10.6), 67(24.3), 64(11.4), 55(10.3), 50(11.0), 44(100.0). 4-Ethoxv-3-methoxyphenvlglvcerol-B-guaiacvl ether (62) *H N M R (CDC13) 8: 1.48(3H, t, J=7Hz), 3.88(6H, two close singlets), 3.91(3H, s), 4.02-4.20(3H, m), 4.99(2H, m), 6.80-7.15(7H, m). MS m/z(%): 348(3.4, M), 300(6.6), 208(10.0), 206(6.6), 195(5.4), 182(16.0), 181(30.8), 180(33.2), 165(15.2), 164(10.3), 153(15.3), 152(21.1), 151(39.2), 150(93.0), 138(35.0), 137(33.7), 125(30.2), 124(78.1), 123(22.3), 122(13.9), 121(20.2), 110(10.0), 109(68.3), 97(10.7), 95(17.5), 93(22.4), 91(13.2), 83(13.3), 81(31.0). 4-O-ethvldehydrodiisoeugenol (64) ' H N M R (CDCI3) 8: 1.39(3H, d, J=7Hz), 1.46(3H, t, J=8Hz), 2.07(3H, dd, J1=7Hz, J 2-0-lHz), 3.43-3.52(lH, m), 3.87(3H, s), 3.90(3H, s), 4.10(2H, q, J=8Hz), 5.12(1H, d, J=9Hz), 6.07-6.17(lH, m), 6.34-6.42(lH, m), 6.77-7.01(5H, m). MS m/z(%): 355(24.1, M+l), 354(100.0, M) , 137(14.1), 91(16.7), 77(21.0), 65(14.6), 55(19.2). M.P.: 121-122°C. 2,3-Dihycho-5-fonnvl-7-meftoxv-3-m (65) 179 ' H N M R (CDCI3) 8: 1.44-1.49(6H, overlap of a doublet and a triplet), 3.53-3.62(lH, m), 3.87(3H, s), 3.96(3H, s), 4.12(2H, q, J=8Hz), 5.27(1H, d, J=9Hz), 6.85-6.98(3H, m, aromatic), 7.35-7.42(2H, aromatic), 9.86(1H, s). MS m/z(%): 343(21.9, M+l), 342(100.0, M) , 314(11.9), 313(21.3), 157(11.4), 149(12.7), 137(33.7), 103(10.3), 95(19.1), 91(29.9), 89(16.0), 79(18.2), 78(17.6), 77(58.2), 76(18.0), 65(33.2), 64(12.5), 63(19.5), 55(31.6), 53(16.1), 52(10.6), 51(28.6), 44(22.0), 43(15.0), 41(12.9), 40(24.3). M.P.: 93-95°C. 5-Carboxv-2,3-m^vdro-7-methoxv-3-methvl-2-(4-ethoxv-3-methoxyphenvl)benzofuran (66) (isolated as its methyl ester) ' H N M R (CDCI3) 8: 1.42(3H, d, J=7Hz), 1.46(3H, t, J=8Hz), 3.49-3.58(lH, m), 3.87(3H, s), 3.91(3H, s), 3.94(3H, s), 4.11(2H, q, J=8Hz), 5.22(1H, d, J=9Hz), 6.84-6.97(3H, aromatic), 7.51(2H, broad s, aromatic). MS m/z(%): 372(18.3, M) , 167(21.0), 149(58.5), 111(10.4), 83(11.9), 77(10.7), 76(10.9), 71(49.4), 70(50.5), 69(18.8), 65(10.9), 59(16.6), 57(100.0), 56(24.7), 55(47.7), 44(32.1), 43(69.4), 42(13.3), 41(61.0), 40(61.2). M.P.: 143-145°C. 5-Carboxy-2,3-dihvdro-2-r(6)-ethvl (l)-methvl 2-(E)-4-(Z)-2,4-hexadienedioate-3-vll-180 7-methoxv-3-methvlbenzofuran (67) (isolated as its methyl ester) COOH 'H NMR (CDClj) 8: 1.27(3H, t, J=8Hz), 1.39(3H, d, J=7Hz), 3.42-3.47(lH, m), 3.69(3H, s), 3.90(3H, s), 3.95(3H, s), 4.17(2H, q, J=8Hz), 5.18(1H, d, J=8Hz), 6.07(1H, d, J=12-13Hz), 6.14(1H, d, J=0-lHz), 6.94(1H, dd, J ias12-13Hz, J2=0-lHz), 7.49-7.54(2H, aromatic). NOE difference spectrum: When irradiated at 5.18 ppm, positive signals appeared at 6.14 and 6.94 ppm. MS m/z(%): 404(23.8), 373(18.3), 358(35.3), 345(17.3), 344(37.7), 331(27.9), 330(27.7), 315(15.3), 314(17.2), 299(40.0), 298(19.6), 285(20.0), 271(27.7), 269(17.1), 267(22.6), 179(100.0), 167(15.5), 151(23.4), 150(22.3), 149(33.3), 123(26.0), 59(17.1), 57(28.5), 41(20.5). Exact mass calcd. for QiH„Oa: 404.1472; found (HRMS): 404.1479. 2.3-Dmvdro-5-formvl-2-r(6)-ethvl (1 )-methvl 2-m)-4-(Z)-2.4-hexadienedioate-3-vl)- 7-methoxv-3-methvlbenzofuran (68) 181 CHO 'H N M R (CDClj) 8: 1.26(3H, t, J=8Hz), 1.41(3H, d, J=8Hz), 3.45-3.54 (IH, m), 3.69(3H, s), 3.96(3H, s), 4.18(2H, q, J=8Hz), 5.23(1H, d, J=8Hz), 6.09 (IH, d, J=12Hz), 6.13(1H, m), 6.91-6.97(1H, m), 7.30(1H, broad s), 7.35(1H, broad s ) , 9.83(1H, s). NOE difference spectrum: When irradiated at 5.23 ppm, positive signal appeared at 6.13 ppm and 6.1-6.97 ppm. MS m/z(%): 374(31.0), 342(25.9), 329(24.8), 328(72.2), 315(29.1), 314(42.0), 301(72.7), 300(44.1), 296(29.3), 270(23.2), 269(100.0), 241(61.3), 240(21.3), 213(54.3), 209(21.1), 181(20.9), 155(40.0), 150(35.9), 149(23.3), 135(20.2), 123(67.4), 115(24.2), 111(26.6), 91(28.8), 79(27.0), 77(37.5), 69(26.8), 65(24.1), 59(69.1), 57(25.8), 55(32.6), 53(21.5), 51(25.1), 44(37.8), 43(51.1), 41(33.9). Exact mass calcd. for C ^ O , : 374.1366; found (HRMS): 374.1369. 2.3-Dihvdro-5-formvl-2-r(6)-ethvl (l)-methvl 2-(Tf)-4-(E)-2,4-hexadienedioate-3-vri- 7-methoxv-3-methvlbenzofuran (69) 'H N M R (CDC1,) 8: 1.33(3H, t, J=8Hz), 1.49(3H, d, J=8Hz), 3.40-3.46(lH, m), 182 CHO C0 2CH 3 69 3.76(3H, s), 3.98(3H, s), 4.16(2H, q, J=8Hz), 5.25(1H, d, J=7Hz), 6.16(1H, d, J=16Hz), 6.16QH, s, broad), 7.33(1H, s, broad), 7.40(1H, s, broad), 8.50(1H, d, broad, J=16Hz), 9.85(1H, s). NOE difference spectrum: When irradiated at 5.25 ppm, positive signal appeared at 6.16 ppm (both the singlet and the doublet). MS m/z(%): 374(28.7), 342(11.8), 328(37.2), 314(22.8), 301(100.0), 300(23.4), 269(61.9), 268(29.9), 241(38.6), 213(26.1), 150(25.3), 123(25.1), 111(27.8), 77(20.2), 59(25.4), 57(20.9), 55(24.1), 44(33.3), 43(33.9), 41(22.0). Exact mass calcd. for QpHjA: 374.1366; found (HRMS): 374.1371. 2-Acetvl-4-formvl-6-methoxvuhenyl 4-ethoxv-3-methoxvbenzoate (70) ' H N M R (CDCI3) 8: 1.53(3H, t, J=8Hz), 2.59(3H, s), 3.92(3H, s), 3.97(3H, s), 4.22(2H, q, J=8Hz), 6.99(1H, d, J=8Hz), 7.68-7.72(2H, m), 7.90(1H, dd, J ^ H z , J2=2Hz), 7.96(1H, d, J=2Hz), 10.03(1H, s). MS m/z(%): 372(2.4), 180(36.8), 179(100.0), 123(37.3), 65(20.7), 59(31.5), 183 57(26.4), 55(22.6), 51(20.1), 43(55.9), 41(36.9), 31(20.7). 4,4'-Diethyldehydrodivanillin (74) : H N M R (CDC13) 8: 1.12(6H, t, J=8Hz), 3.98(6H, s), 4.02(4H, q, J=8Hz), 7.46(2H, d, J=l-2Hz), 7.51(2H, d, J=l-2Hz), 9.91(2H, s). MS m/z(%): 359(10.4, M+l), 358(52.2, M), 302(20.8), 241(10.7), 91(16.2), 32(16.9), 28(100.0). M.P.: 142-144°C. 4,4,-Diethoxv-3,3/-dimethoxv-5-carboxv-5/-formvlbiphenyl (75) (isolated as its methyl ester) ' H N M R (CDCI3) 8: 1.09(3H, t, J=7 Hz), 1.12(3H, t, J=7 Hz), 3.91(3H, s), 3.96(3H, s), 3.97(3H, s), 3.94-4.04(4H, m), 7.43(1H, d, J=l-2 Hz), 7.49(1H, d, J=l-2 Hz), 7.65(2H, s, broad), 9.90(1H, s). MS m/z (%): 389(M+1, 22.5), 388(M, 100.0), 360(20.7), 332(18.6), 301(15.1), 300(56.9), 272(15.5), 271(15.8). j Exact mass calcd. for C^H^O, : 388.1523; found (HRMS): 388.1529. M.P.: 110-112°C. 184 4,4/-Diethvldehvdrodivanillic acid (76) (isolated as its dimethyl ester) J H N M R (CDCI3) 8: 1.10(6H, t, J=7 Hz), 3.90(6H, s), 3.95(6H, s), 3.96(4H, q, J=7Hz), 7.62(4H, s). MS m/z (%): 419(M+1, 22.8), 418(M, 100.0), 390(22.6), 358(21.2), 330(82.6), 313(26.8), 299(20.5), 284(21.6), 242(29.3), 209(21.3), 59(25.3), 57(20.7), 43(21.5), 41(22.9). Exact mass calcd. for C ^ O s : 418.1628 ; found(HRMS): 418.1632. M.P.: 102-104°C. (l)-Ethyl (6)-methyl 2-(2-ethoxv-3-methoxv-5-formvlphenyl)-4-carboxv-2.4- hexadienedioate (77) (isolated as its methyl ester) ' H N M R (CDCI3) 8: 1.21(3H, t, J=7 Hz), 1.27(3H, t, J=7 Hz) 3.76(3H, s), 3.82(3H, s), 3.94(3H, s), 4.10(2H, q, J=7 Hz), 4.16(2H, q, J=7 Hz), 6.80(1H, d, J=l-2 Hz), 7.26(1H, d, J=l-2 Hz), 7.50(1H, d, J=l-2 Hz), 7.54(1H, d, J=l-2 Hz), 9.93(1H, s). MS m/z (%): 420(M, 4.6), 361(5.6), 347(14.8), 315(7.1), 287(27.7), 259(13.5), 149(12.7), 115(18.1), 91(14.2), 77(20.8), 69(13.6), 63(13.9), 57(17.3), 55(20.1), 51(18.7), 50(15.0), 45(19.), 44(100.0). 185 Exact mass calcd. for G A C v 420.1421; found(HRMS): 420.1429. (1 )-Ethyl (6)-methvl 2-(2-ethoxv-3-methoxv-5-carboxyphenvl)-4-carboxv-2.4- hexadienedioate (78) (isolated as its dimethyl ester) 'H NMR (CDC13) 8: 1.21(3H, t, J=7Hz), 1.25(3H, t, J=7Hz), 3.76(3H, s), 3.80(3H, s), 3.92(6H, s), 4.05(2H, q, J=7Hz), 4.14(2H, q, J=7Hz), 6.77(1H, d, J=l-2Hz), 7.24(1H, d, J=l-2Hz), 7.63(1H, d, J=l-2Hz), 7.73(1H, d, J=l-2Hz). MS m/z (%): 450(M, 18.2), 391(46.3), 377(58.5), 359(21.2), 345(48.1), 331(40.8), 318(23.7), 317(100.0), 303(39.2), 287(23.0), 285(34.9), 273(24.7), 262(42.6), 259(26.8), 258(31.9), 257(22.0), 234(33.3), 233(22.9), 203(67.2), 175(20.5), 149(42.0), 115(25.6), 91(31.5), 77(32.4), 71(20.9), 69(25.2). Exact mass calcd. for C^llJD^: 450.1526; found(HRMS): 450.1524. Pyrene-1,6-dione (80) !H NMR (DMSO-(L., 8): 6.72 (2H, d, J=10 Hz), 8.02(2H, d, J=10Hz), 8.14(2H, d, J=8Hz), 8.42(2H, d, J=8Hz). MS m/z (%): 234(M+2, 14.3), 233(M+1, 19.3), 232(M, 100.0), 204(42.8), 176(89.0), 92(30.9), 91(62.4), 88(79.5), 87(23.0), 75(34.6). 186 Exact, mass calcd. for C 1 6 r L 0 2 : 232.0540; found (HRMS): 232.0523. M.P.: sublimes at around 240°C. Pyrene-1.8-dione (81) ' H N M R (DMSO-cL,, 8): 6.69(2H, d, J=9Hz), 7.99-8.03(4H, overlap of a doublet with a singlet at 8.02), 8.54(2H, s). MS m/z (%): 234(M+2, 27.0), 233(M+1, 20.5), 232(M, 100.0), 205(21.9), 204(44.5), 176(95.9), 175(20.0), 150(20.4), 92(26.7), 91(52.1), 88(82.5), 87(24.2), 75(38.7). Exact mass calcd. for C 1 6 H 8 0 2 : 232.0540; found (HRMS): 232.0519. M.P.: sublimes at around 190°C. 2,6-Dichloro-p-benzoquinone (83) ! H N M R (CDC13) 8: 7.05 (s) MS m/z (%): 180(10.9), 178(51.8), 176(68.1), 150(13.4), 148(21.8), 141(16.8), 122(36.7), 120(56.8), 116(13.0), 115(15.6), 113(49.1), 90(26.8), 88(77.7), 87(23.0), 85(31.5), 62(47.8), 61(23.0), 60(100.0), 53(94.3), 52(12.1), 50(47.9), 49(38.9). 187 Exact mass calcd. for C 6 H 2 0 2 C1 2 (CI 3 5 isotope): 175.9432; found (HRMS): 175.9428. 188 References for chapter 3 1. K . Lundquist, and T.K. Kirk, Phytochem. 17, 1676, 1978. 2. M . Shimada, F. Nakatsubo, T.K. Kirk, and T. Higuchi, Arch. Microbiol. 129, 321, 1981. 3. T.K. Kirk, and R.L. Farrell, Annu. Rev. Microbiol. 41, 465, 1987. 4. M.S.A. Leisola, D.C. Ulmer, R. Waldner, and A . Fiechter, J. Biotechnol. 1, 331, 1984. 5. D.C. Ulmer, M.S.A. Leisola, and A . Fiechter, J. Biotechnol. 1, 13, 1984. 6. S.D. Haemmerli, M.S.A. Leisola, D. Sanglard, and A . Fiechter, J. Biol. Chem. 261, 6900, 1986. 7. T. Tonon, and E. Odier, Appl. Environ. Microbiol. 54, 466, 1988. 8. P.J. Harvey, H.E. Schoemaker, and J .M. Palmer, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 145, 1987. 9. H . Wariishi, and M . H . Gold, FEBS Letters, 243, 165, 1989. 10. B.D. Faison, T.K. Kirk, and R.L. Farrell, Appl. Environ. Microbiol. 52, 251, 1986. 11. M.S.A. Leisola, B . Schmidt, U . Thaner-Wyss, and A . Fiechter, FEBS Letters, 189, 267, 1987. 12. M.S.A. Leisola, S.D. Haemmerli, R. Waldner, H.E. Schoemaker, H.W.H. Schmidt, and A . Fiechter, Cellulose Chem. Technol. 22, 267, 1988. 13. M . Shimada, T. Hattori, T. Umezawa, T. Higuchi, and T. Okamoto, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 151, 1987. 189 14. M . Shimada, T. Hattori, T. Umezawa, T. Higuchi, and K . Uzura, FEBS Letters, 221, 327, 1987. 15. M.S.A. Leisola, S.D. Haemmerli, J.D.G. Smit, J. Trailer, R. Waldner, H.E. Schoemaker, and H . Schmidt, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 81, 1987. 16. T. Hattori, M . Shimada, T. Umezawa, T. Higuchi, M.S.A. Leisola, and A . Fiechter, Agric. Biol . Chem. 52, 879, 1988. 17. H.W.H. Schmidt, S.D. Haemmerli, H.E. Schoemaker, and M.S.A. Leisola, Biochem. 28, 1776, 1989. 18. T.K. Kirk, M . Tien, P.J. Kersten, M.D. Mozuch, and B . Kalyanaraman, Biochem. J. 236, 279, 1986. 19. J.S. Dordick, M . A . Marietta, and A . M . Klibanov, Proc. Natl. Acad. Sci. USA, 83, 6555, 1986. 20. N.G. Lewis, R.A. Razal, and E. Yamamoto, Proc. Nad. Acad. Sci. USA, 84, 7925, 1987. 21. F. Cui, and D. Dolphin, in Plant Cell Wall Polymers: Biogenesis and Biodegradation. Ed. N.G. Lewis, and M . G . Paice, Am. Chem. Soc. Washington DC, 1989, chp. 37. 22. A . Sakakibara, in Recent Advances in Lignin Biodegradation Research, Ed. T. Higuchi, H . - M . Chang, and T.K. Kirk, Uni. Publishers, Tokyo, 1983, pl2. 23. M . Tien, and T.K. Kirk, Proc. Natl. Acad. Sci. USA, 81, 2280, 1984. 24. V . Renganathan, K , Mike, and M . H . Gold, Arch. Biochem. Biophys. 241, 304, 1985. 25. V . Renganathan, K . Mike, and M . H . Gold, Arch. Biochem. Biophys. 246, 155, 1986. 190 26. H . Numba, F. Nakatsubo, and T. Higuchi, Wood Res. 69, 52, 1986. 27. J.K. Glenn, M . A . Morgan, M . B . Mayfield, M . Kuwahara, and M . H . Gold, Biochem. Biophys. Res. Commun. 114, 1077, 1983. 28. M . H . Gold, M . Kuwahara, A . A . Chiu, and J.K. Glenn, Arch. Biochem. Biophys. 234, 353, 1984. 29. T. Umezawa, F. Nakatsubo, and T. Higuchi, Agric. Biol. Chem. 47, 2677, 1983. 30. T. Higuchi, Wood Res. 67, 47, 1987. 31. T. Katagama, and T. Higuchi, Abst. 25th Symp. Lignin Chem. Fukuoko, Japan, 1980, p5. 32. F. Nakatsubo, T.K. Kirk, M . Shimada, and T. Higuchi, Arch. Microbiol. 128, 416, 1981. 33. L . A . Anderson, V . Renganathan, A . A . Chiu, T .M. Lohr, and M . H . Gold, J. Biol. Chem. 260, 6080, 1985. 34. B.W. Griffin, J.A. Peterson, and R.W. Estabrook, in The Porphyrins, Ed. D. Dolphin, Academic Press, New York, 1979, Vol . VII, p333. 35. V . Renganathan, K . Miki , and M . H . Gold, Biochem. 26, 5127, 1987. 36. J.A. Bumpus, M . Tien, D. Wright, and S.D. Aust, Science, 228, 434, 1985. 37. M . Shimada, T. Habe, T. Umezawa, T. Higuchi, and T. Okamoto, Biochem. Biophys. Res. Commun. 122, 1247, 1984. 38. T. Habe, M . Shimada, and T. Higuchi, Mokuzai Gakkaishi, 31, 54, 1985. 39. E. Adler, Wood Sci. Technol. 11, 169, 1977. 40. K . M i k i , V . Renganathan, and M . H . Gold, Biochem. 25, 4790, 1986. 41. T. Umezawa, F. Nakatsubo, and T. Higuchi, Arch. Microbiol. 131, 124, 1982. 42. F. Nakatsubo, T.K. Kirk, M . Shimada, and T. Higuchi, Arch. Microbiol. 126, 416, 1981. 191 43. T. Umezawa, M . Shimada, T. Higuchi, and K . Kusai, FEBS Letters, 205, 287, 1986. 44. T. Umezawa, and T. Higuchi, FEBS Letters, 205, 293, 1986. 45. K . Mik i , V . Renganathan, M . B . Mayfield, and M . H . Gold, FEBS Letters, 210, 199, 1987. 46. T. Umezawa, and T. Higuchi, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 63, 1987. 47. K . Mik i , V . Renganathan, M . B . Mayfield, and M . H . Gold, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 69, 1987. 48. P.J. Kersten, M . Tien, B . Kalayanaraman, and T.K. Kirk, J. Biol. Chem. 260, 2609, 1985. 49. T.K. Kirk, J .M. Harkin, and E.B. Cowling, Biochim. Biophys. Acta, 165, 134, .1968. 50. Y . Kamaya, and T. Higuchi, Mokuzai Gakkaishi, 29, 789, 1983. 51. K. Krisnangkura, and M . H . Gold, Holzforschung, 33, 174, 1979. 52. T. Fukuzumi, in Lignin Biodegradation: Microbiology, Chemistry, and Potential Applications. Ed. by T.K. Kirk, T. Higuchi, and H . - M . Chang, CRC Press, 1980, p73. 53. T. Umezawa, and T. Higuchi, Abst. of papers, 27th symposium on lignin chemistry, Nagoya, Japan, 1982, p l O l . 54. J. Jokela, J. Pellinen, and M . Salkinoja-salonen, Appl. Environ. Microbiol. 53, 2642, 1987. 55. J. Jokela, J. Pellinen, M . Salkinaja, and G. Brunnow, Appl. Microbiol. Biotechnol. 23, 38, 1985. 56. Y . Kamaya, and T. Higuchi, Wood Res. 70, 25, 1984. 192 57. H.E. Schoemaker, E . M . Meijer, M.S.A. Leisola, S.D. Haemmerli, R. Waldner, D. Sanglard, and H.W.H. Schmidt, in Plant Cell Wall Polymers: Biogenesis and Biodegradation. Ed. by N.G. Lewis, and M . G . Paice, Am. Chem. Soc. Washington DC, 1989, chp. 33. 58. T. Umezawa, S. Kawai, S. Yokota, and T. Higuchi, Wood Res. 73, 8, 1986. 59. K . E . Hammel, B. Kalyanaraman, and T.K. Kirk, J. Biol . Chem. 261, 16948, 1986. 60. D. Sanglard, M.S.A. Leisola, and A . Fiechter, Enzyme Microb. Technol. 8, 209, 1986. 61. K . E . Hammel, B . Kalyanaraman, and T.K. Kirk, Collq. INRA 40 (Lignin Enzyme Microb. Degrad.), 45, 1987. 62. T. Katayama, F. Nakatsubo, and T. Higuchi, Mokuzai Gakkaishi, 27, 223, 1981. 63. F. Nakatsubo, and T. Higuchi, Holzforschung, 29, 193, 1975. 64. B. Leopold, Acta Chem. Scand. 4, 1523, 1950. 65. K. Elbs, and H . Lerch, J. Prakt. Chemie. 93, 1, 1916. 66. T. Umezawa, T. Higuchi, and F. Nakatsubo, Agric. Biol. Chem. 47, 2945, 1983. 67. L.F. Fieser, and M . Fieser in Reagents for Organic Synthesis, John Wiley and Sons, 1967, pl91. 68. S.D. Haemmerli, H.E. Schoemaker, H.W.H. Schmidt, and M.S.A. Leisola, FEBS Letters, 220, 149, 1987. 69. Beilsteins Handbuch der Organischen Chemie, Springer-Verlag, Berlin, 6, 964b. 70. Beilsteins Handbuch der Organischen Chemie, Springer-Verlag, Berlin, 8, 256b. 71. Beilsteins Handbuch der Organischen Chemie, Springer-Verlag, Berlin, 8 I, 618h. 72. Beilsteins Handbuch der Organischen Chemie, Springer-Verlag, Berlin, 6 III, 6333e. 193 Chapter 4 Redox mediation in lignin biodegradation Lignin is a three-dimensional, water-insoluble high molecular weight polymer and the exact pathway by which it is metabolised by fungi is still unknown. Although lignin degrading enzymes, lignin peroxidase (LiP) and manganese-dependent peroxidase (MnP) have been isolated1,2 and partially characterized by spectroscopic and other physical methods,3-11 the enzymes are successful only in cleaving the C-C and C-O bonds of low molecular weight lignin model compounds. Effective depolymerization or degradation of isolated lignin polymers by purified enzyme or enzyme combination has not been achieved. It is evident, however, that lignin peroxidase alone can at least modify (oxidize) lignin 1 2 , 1 3 and in some cases release low molecular weight fragments.1,2 It is interesting and also important to understand how an enzyme (with a molecular weight over 40,000) interacts with an irregular, cross-linked, water-insoluble lignin polymer. Due to the physical properties and complex chemical structure of lignin, it is unlikely that all the substructures of lignin can approach and fit into the active site of lignin degrading enzymes. Both L iP and MnP oxidize lignin model compounds by single electron oxidation.14 Redox mediators which function as electron carriers between the enzyme and the lignin substrate have been suggested to be involved in lignin biodegradation. 194 4.1 Manganese complexes as mediators in lignin biodegradation The manganese dependent peroxidase (MnP) is another enzyme isolated from the ligninolytic culture of P. chrysosporium. The reactions catalyzed by this enzyme7"11 have not been extensively studied and its role in lignin degradation is still not clear. MnP was first found to be dependent on Mn(H) and a-hydroxycarboxylic acids for its activity.7 A l l the enzyme activity was lost after dialysis but could be rapidly restored upon addition of Mn(II), suggesting Mn(TJ) was loosely bound to the enzyme.8 It was found later15 that MnP could oxidize Mn(II) to give Mn(ffl) in the presence of hydrogen peroxide and several a-hydroxycarboxylic acids, and the Mn(ni)-oc-hydroxycarboxylic acid complex could in turn oxidize the L iP substrates while being reduced to the Mn(U) complex. The function of the a-hydroxycarboxylic acids was to stabilize Mn(III). The manganese complexes, therefore, are the mediators for MnP catalyzed oxidation of lignin model compounds. L iP and MnP oxidize lignin model compounds by single-electron oxidations. Electrochemical methods, which can initiate one-electron oxidation processes at the anode, have been used to study the oxidation of lignin model compounds.16,17 Electrochemical oxidation of Mn(II) to give Mn(UI) has also been reported.18,19 In this study, the mediating ability of the manganese complexes was studied in an enzyme free system in which electrochemical and biomimetic methods were used to generate Mn(IU) complexes. Mew-tetra(2,6-dichloro-3-sulfonatophenyl)porphyrin iron chloride (TDCSPPFeCl) was used as a biomimetic catalyst to mimic the function of MnP. A polymer dye, Poly B-411 (84), was used as an oxidizable lignin model compound. It has been shown that Poly B-411 is a convenient water-soluble lignin model and its 195 0 NH NH S0 3Na * | f J S0 3Na 0 NH 2 84 n decolorization has been correlated with Ugnin degradation.8,20 The decolorization of Poly B-411 can be easily quantified by foUowing its absorption at 596 nm. The reason that the mediating ability of manganese complexes is studied in an enzyme free system is that separate evidence would be provided to support its function. Secondly, any mediating power of the manganese complexes in the model system would indicate that the oxidized complexes could function without association with the enzyme. 4.1.1 Results 4.1.1.1 Cyclic voltammetry (CV) of manganese complexes Cyclic voltammograms (CV) of several manganese(II) species were obtained at pH 4.5 in the presence of several ligands. CV of manganese sulfate (10 mM) containing 50 mM lactic acid exhibited two oxidation peaks at 0.82 V and 1.37 V, respectively. The oxidation peak at 1.37 V disappeared when the CV was carried out in the presence of a large excess of lactic acid. The CV of manganese sulfate alone also showed a peak at 1.37 V. The oxidation peak at 0.82 V was therefore due to the Mn-lactate complex and that at 1.37 V to "uncomplexed" manganese ions. The CV of manganese sulfate in the presence of various carboxylic acids and inorganic ligands was 196 Table 4.1 Cyclic voltammetry of manganese sulfate (10 mM solution, pH 4.5) in the presence of various ligands*3 L- DL-Ligand F S042 C104" CH3C02" Lactate Malate Malonate Pyruvate Citrate D-tartrate H20 concen- >100 100 100 200 70 60 10 50 10 10 tration (mM) oxidation 0.56" 0.95b) 0.93b) 0.85b> 0.82b) 0.91b) 0.8 l b ) 0.92c> 0.83c) 0.86c) 1.37c) potential (V) *} sweep rate 50 mv/second,b) pseudo-reversible, c ) irreversible similarly studied and the results are summarized in Table 4.1. Manganese formed complexes with DL-malate, malonate, pyruvate, citrate, L-lactate, and D-tartarate. Manganese seemed even to be complexed by inorganic ligands since oxidation peaks at lower potentials than 1.37 V were observed in the presence of these ligands. Manganese dioxide, however, was formed on the surface of the working electrode after several runs when the inorganic anions were used as ligands, indicating the relative instability of the complexes having these inorganic ligands. Manganese dioxide was also formed on the electrode when the carboxylic acids were present at lower concentrations than those listed in Table 4.1. x To confirm that the manganese(JJ.)-complexes was oxidized to Mn(m) during the electrochemical oxidation, the manganese-malonate complex in its higher oxidation state was prepared by controlled potential oxidation. The Mn(in)-malonate generated by controlled potential oxidation and that made chemically by mixing manganese(HI) acetate and malonic acid gave the same UV-vis spectra (k^ 270 nm). Addition of sodium pyrophosphate to a solution of electrochemically generated Mji(rfl)-malonate gave Mn(m)-pyrophosphate, which has a characteristic absorption at 296 nm. Mixing manganese(III) acetate and malic acid in glacial acetic acid gave a light brown gelatinous precipitate. The precipitate was soluble in pH 4.5 buffer (containing malic acid) and the aqueous solution showed a similar UV-vis spectrum as that of manganese(HI)-malate prepared by mixing manganese(IU) acetate and malic acid in pH 4.5 acetate buffer, suggesting that the precipitate was Mn(IU)-malate. Addition of the Mn(HI)-malate powder to a solution containing Poly B-411 caused instant decolorization of the dye. Efforts to recrystallize the Mn(IIT)-malate precipitate failed. Elemental analysis of the Mn(JJJ.)-malate supports a formula of M r ^ Q H A V S r L O . 198 4.1.1.2 Manganese mediated decolorization of Poly B-411 The CV of 0.02% Poly B-411 in pH 4.5 acetate buffer showed an oxidation peak at 0.75 V. The controlled potential oxidation (decolorization) of Poly B-411 was, however, very slow, probably because of its polymeric nature. When manganese sulfate (added to a final concentration of 0.1 mM which was the optimum concentration for MnP8) and a-hydroxycarboxylic acids or related compounds were added to the solution, the electrochemical decolorization of Poly B-411 was accelerated. Figure 4.1 shows the change of the absorbance of the dye at 596 nm with oxidation time in the presence of different amounts of lactic acid at a constant concentration of manganese sulfate (0.1 mM). Maximum acceleration was achieved when the concentration of lactic acid was Table 4.2 Comparison of the optimum concentrations of carboxylic acids in the electrochemical decolorization of Poly B-411 and those in the MnP enzyme system. Ligand Optimum concentration Concentration with (mM) highest current efficiency (mM) Electrochemical Enzyme system system8 (L)-lactic 50-60 50 70 acid malic acid 10 10 10 malonic acid 50 - 60 citric acid 10 - a) (D)-tartaric acid 20 25 a) The current efficiency is higher when the concentration is either 5 or 20 mM but the rate is slower. 199 1.40 Time (min) 0 10 20 30 40 50 60 Figure 4.1 The electrochemical decolorization of Poly B-411 in 0.1 M acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and lactic acid at various concentrations (mM), 1, 0; 2, 10; 3, 20; 4, 30; 5, 50; 6, 60; 7, 70; 200 50 or 60 m M (Figure 4.1). The decolorization of Poly B-411 in the presence of manganese sulfate and various amounts of D-tartaric acid, citric acid, malic acid, and malonic acid are shown in Figures 4.2-4.5. The rate of decolorization of Poly B-411 depended on the concentration of the carboxylic acids and the optimum concentrations for maximum decolorization are summarized in Table 4.2. The decolorization of Poly B-411 in the presence of manganese sulfate and various carboxylic acids at their optimum concentration is shown in Figure 4.6 As manganese sulfate gave a pseudo-reversible cyclic voltamogram in the presence of 0.1 M sodium perchlorate, the effect of manganese and perchlorate on the decolorization of Poly B-411 was studied. Dye decolorization was accelerated by addition of 0.1 m M manganese sulfate and 0.1 M sodium perchlorate, but the current efficiency was lower than with the carboxylic acid ligands and manganese dioxide was found on the working electrode surface. Very small acceleration and low current efficiency were observed with pyruvic acid. To confirm that Mn(UI) complexes were the actual oxidant of Poly B-411, a dye solution was added to the anode compartment of a electrochemical cell and the glassy carbon anode was placed in a dialysis bag. Manganese sulfate (0.1 mM) and lactic acid were added to both sides of the dialysis bag. Decolorization of the dye solution was observed after 2 hours oxidation at 0.8 V . No dye decolorization was observed after 5 hours oxidation in a control experiment lacking manganese sulfate and lactic acid. The decolorization of Poly B-411 by TDCSPPFeCl and mCPBA in the presence or absence of manganese-lactate is shown in Figure 4.7. The decolorization was more extensive and faster in the presence of manganese-lactate. 201 0.0 10,0 20,0 30,0 40,0 50,0 60,0 T i n e (min) Figure 4.2 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of malic acid (mM), 1, 10; 2, 15; 3, 5; 202 A A596 2.20 2.10 -2.00 1.90 -1.80 -1.70 1.60 1.50 V \ \ N N V \ N 1 1 1 1 1 1 — 0,0 10.0 20.0 30.0 40.0 50.0 60.0 Tine (min) Figure 4.3 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of (D)-tartaric acid (mM), 1, 10; 2, 20; 3, 25; 203 A596. 0 10 20 30 40 50 60 Time (min) ;ure 4.4 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of citric acid (mM), 1, 10; 2, 20; 3, 5; Figure 4.5 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various concentrations of malonic acid (mM), 1, 10; 2, 50; 3, 60; 4, 40; A596 to o ON 2.20 2.10 • 2.00 1.90 1.80 1.70 1.60 -1.50 ' 1.40 ~i 1.30 " T(min) o 10 20 30 40 50 60 Figure 4.6 The electrochemical decolorization of Poly B-411 (0.02%) in 0.1 M pH 4.5 acetate buffer at room temperature in the presence of manganese sulfate (0.1 mM) and various carboxylic acids at their optimum concentrations, 1, no ligand; 2, 5 m M citric acid; 3, 20 m M (D)-tartaric acid; 4, 60 mM (L)-lactic acid; 5, 10 mM malic acid; 6, 50 m M malonic acid; 0.70-Figure 4.7 i r 6 7 Time (min) The decolorization of Poly B-411 (0.005% in 25 mL 0.1 M pH 4.5 acetate buffer) by TDCSPPFeCl (1x10* mole) and mCPBA (2.5X10-5 mole) in the presence of manganese sulfate (2.5x105 mole) and lactic acid (1.5x10^ mole), 1, without TDCSPPFeCl; 2, without manganese sulfate; 3, complete reaction; 4.1.2 Discussion Mn(IU) has a standard reduction potential of 1.51 V and is stabilized in aqueous solution at very low pH or by formation of coordination complexes. The carboxylic acids used in this work form relatively weak Mn(JH) complexes of reasonable oxidizing power, unlike some other strong complexing agents21 which dramatically decrease the oxidizing power of Mn(III). Numerous complexes between Mn(IU) and a-hydroxycarboxylic acids have been reported.22"24 The complexes of Mn(HI) with simple a-hydroxycarboxylic acids, on the other hand, are not well known. Hydroxyhexafluoroisobutyric acid [HjfHHIB)] gives a stable Mn(III) complex [Mn(HFflB)3]3" which may be crystallized as its caesium complex. H2(HHIB), however, is a non-oxidizable ligand and other simple non-oxidizable carboxylic acids such as acetic acid also form stable Mn(III) compounds. Our inability to characterize the complex between malate and Mn(III) probably results from the redox chemistry of the ligand together with the probability of oligomer formation due to the bidentate nature of the ligand. Nevertheless, a-hydroxycarboxylic acids and related acids clearly stabilize Mn(IJJ) and inhibit its disproportionation which otherwise results in the precipitation of manganese dioxide. We have shown the mediating power of a readily diffusible Mn(UI)-a-hydroxycarboxylic acid complex by generating the Mn(III) electrochemically in the presence of lactate and the subsequent oxidation of Poly B-411 separated from the electrode by a semipermeable membrane. Table 4.1 and Figure 4.6 show that the efficiency of the manganese complexes as redox mediators are not direcdy related to their redox potentials. Rather, the stability of the manganese(III) complexes and their rates of electron transfer, which are related 208 to their structures, will be key factors in their roles as effective mediators. The optimum concentration of lactic acid and malic acid (Table 4.2) in the experiments described above are about the same as that of the enzyme system,8 suggesting that the electrochemical system used does indeed mimic and describe the enzymic system. Additional evidence is provided that manganese complexes work as diffusible redox mediators for lignin model compound oxidation. The mediating power of manganese complexes are not just restricted to within the MnP enzyme system. The biomimetic oxidation of Poly B-411, a lignin model compound, by electrochemical and metalloporphyrin/oxidant systems are also accelerated by the presence of manganese complexes. While malonic acid was found to be one of the most effective ligand for the formation of manganese complexes which exhibit maximal mediating power, it was not clear that nature made use of this important cellular metabolite during wood degradation by P. chrysosporium. However, after completing this work, we learned that Gold et aP5 have recently found malonic acid could significantly stimulate MnP activity. The effective decolorization of Poly B-411 by TDCSPPFeCl and mCPBA in the presence of manganese-lactate indicates that TDCSPPFeCl is also able to mimic the function of MnP. mCPBA, a two-electron oxidant, cannot initiate the one-electron oxidation of Poly B-411 (curve 1, Figure 4.7). Addition of TDCSPPFeCl as a biomimetic catalyst causes a rapid but inefficient decolorization (curve 2, Figure 4.7). The TDCSPPFeCl catalyst is rapidly destroyed since it cannot rapidly transfer its oxidizing equivalents readily to the polymeric dye molecule. With the help of manganese-lactate as a mediator, TDCSPPFeCl is able to oxidize Poly B-411 rapidly and more extensively (curve 3, Figure 4.7). TDCSPPFeCl is also more stable in the presence of manganese-lactate since the oxidizing equivalents of oxidized TDCSPPFeCl 209 are rapidly transferred to the substrate by the manganese mediator. 4.2 Veratryl alcohol as a possible mediator in lignin biodegradation Veratryl alcohol is a secondary metabolite of lignin degradation by P. chrysosporium26*1 and plays an important role in lignin biodegradation.3 It can induce the biosynthesis of lignin peroxidase by P. chrysosporium?* It can also stimulate lignin degradation29,30 and benzo(a)pyrene31 oxidation with L iP by protecting the enzyme from inactivation.32"34 Harvey et aP5 found that the presence of a catalytic amount of veratryl alcohol could stimulate the oxidation of anisyl alcohol by L iP and therefore proposed that veratryl alcohol could work as a mediator in lignin biodegradation. Direct evidence for the mediation, however, has not been obtained and more importantly the cation radical of veratryl alcohol, a necessary intermediate in redox mediation, has not been observed spectrally. Enzymic reactions are the most efficient and also the most complex of chemical reactions. In the experiment of Harvey et aP5, veratryl alcohol might stimulate the reaction in conjunction with the enzyme rather than working as a diffusible redox mediator. It is believed that if lignin or its model compounds could be oxidized by a biomimetic system, free of the enzyme, and the reaction could be stimulated by the presence of veratryl alcohol, the mediation role of veratryl alcohol could be established more conclusively in the biomimetic system than in the enzymatic system. In this work, the effect of veratryl alcohol on the oxidation of anisyl alcohol by TDCSPPFeCl and mCPBA and on the electrochemical oxidation of Poly B-411 are studied. The mediation roles of veratryl alcohol are clarified. 210 4.2.1 Results and discussion H a r v e y et a / 3 5 r e p o r t e d t h a t t h e p r e s e n c e o f c a t a l y t i c a m o u n t s o f v e r a t r y l a l c o h o l o r 1 , 4 - d i m e t h o x y b e n z e n e c o u l d s t i m u l a t e t h e o x i d a t i o n o f a n i s y l a l c o h o l a n d 4 -m e t h o x y m a n d e l i c a c i d b y l i g n i n p e r o x i d a s e . T h e o x i d a t i o n o f a n i s y l a l c o h o l b y T D C S P P F e C l a n d m C P B A i n t h e p r e s e n c e o f v a r y i n g a m o u n t s o f v e r a t r y l a l c o h o l i s s h o w n i n F i g u r e 4 . 8 . I n s t e a d o f s t i m u l a t i n g t h e o x i d a t i o n , t h e p r e s e n c e o f v e r a t r y l 0.0 T r 1 10.0 20 .0 30 .0 Veratryl a lcoho l ( pmo les ) F i g u r e 4 . 8 T h e o x i d a t i o n o f a n i s y l a l c o h o l ( 3 x 1 0 s m o l e i n 6 m L p H 3 0 . 1 M p h o s p h a t e b u f f e r ) b y T D C S P P F e C l ( 5 x 1 0 * m o l e ) a n d m C P B A ( 3 x l 0 ' 5 m o l e ) i n t h e p r e s e n c e o f v e r a t r y l a l c o h o l 2 1 1 alcohol inhibited the reaction and the yield of anisaldehyde was the highest in the absence of veratryl alcohol. This observation was not surprising since veratryl alcohol, which had a lower redox potential, could be oxidized more readily than anisyl alcohol. Veratryl alcohol inhibited the reaction by competing with anisyl alcohol for oxidant Similar results were obtained for the oxidation of anisyl alcohol in the presence of 1,4-dimethoxybenzene as shown in Figure 4.9. cu o 'c o •S 10 -33 >-30 -20 ~ 0 0.0 T 1 r 10.0 20.0 30.0 1,4-Dimethoxybenzene (/xmoles) Figure 4.9 The oxidation of anisyl alcohol (3x10* mole in 6 mL 0.1 M pH 3 phosphate buffer) by TDCSPPFeCl (5x10* mole) and mCPBA (3x10* mole) in the presence of 1,4-dimethoxybenzene In the above experiments, the biomimetic catalyst, TDCSPPFeCl, must have a reaction centre more exposed than the enzyme active site. Moreover, both veratryl 212 alcohol and the substrate anisyl alcohol are small molecules. The mediating ability of veratryl alcohol, i f any, could not be easily observed in these model systems. The possibility that veratryl alcohol works as a mediator cannot be excluded based on the above results. As the steric interaction of lignin with Ugnin peroxidase may be quite difficult, a small mediating effect of veratryl alcohol might be quite important for in vivo lignin degradation. Electrochemical methods, which can initiate one-electron oxidation at the electrode, have been used to oxidize Ugnin model compounds.16,17 Unlike other biomimetic systems such as iron porphyrins (hemins) or inorganic one-electron oxidants, the electrode can be considered as 'macromolecular' since the electrode has a limited surface area and its interaction with polymeric molecules, somewhat like the interaction of lignin and lignin peroxidase, is sterically difficult. This property can be utilized to study the mediating function of smaU molecules having mediating potentials. Being polymeric, Poly B-411 is an excellent lignin model for this study. The electrochemical decolorization of Poly B-411 at 0.9 V in the presence of varying amounts of veratryl alcohol is shown in Figure 4.10. Veratryl alcohol was able to stimulate the decolorization significantly in all cases. When the concentration of veratryl alcohol was 0.1 mM, the rate of decolorization remained approximately unchanged over a period of 60 minutes. At higher veratryl alcohol concentrations, 1 and 10 mM, an 'induction period' was needed for maximal decolorization to occur. The higher the concentration, the longer the induction period. During the 'induction period', the absorbance of veratraldehyde at 310 nm rapidly reached a limit but very little Poly B-411 decolorization was observed. The steady decolorization rate after the 'induction period' was slightly higher when the original concentration of veratryl alcohol was higher. One possible explanation is that some oxidation or oxidative 213 0.420 0 10 20 30 40 50 60 Time (min) Figure 4.10 The electrochemical decolorization of Poly B-411 (0.005%) in 0.1 M pH 3 phosphate buffer at room temperature in the presence of various amounts of veratryl alcohol (mM). 1, 0; 2, 0.1; 3, 1; 4, 10. polymerization products of veratryl alcohol might be able to mediate the dye decolorization. A small amount of dimerized product was obtained in the electrochemical oxidation of 4-methoxybenzyl alcohol in acetonitrile.36 The electrochemical decolorization of Poly B-411 was also carried out in the presence of 1 m M veratraldehyde instead of veratryl alcohol. No mediation effect was observed, suggesting that veratryl alcohol rather than veratraldehyde was the mediator. 214 In the above experiments in aqueous solution, Poly B-411 molecules can still approach the electrode and thus be oxidized direcdy. It is assumed that if the polymeric dye were separated from the electrode but were still oxidized in the presence of veratryl alcohol, the decolorization then must be mediated by veratryl alcohol. Poly B-411 was deposited onto a piece of filter paper which was attached direcdy to the glassy carbon anode. Methylene chloride, in which Poly B-411 is insoluble, was used as the solvent. After six hours controlled potential oxidation at 1.2 V , the side of the filter paper which direcdy face the anode was decolorized. The color of the anolyte changed to yellow after the oxidation, suggesting veratryl alcohol was also oxidized. The filter paper with Poly B-411 was not decolorized in a control experiment with no veratryl alcohol in the anolyte solution. It can be concluded from these experiments that veratryl alcohol can indeed work as a diffusible redox mediator for Poly B-411 oxidation. It, however, cannot mediate the oxidation of anisyl alcohol by TDCSPPFeCl and mCPBA. The stimulation effect of veratryl alcohol for the oxidation of anisyl alcohol and 4-methoxymandelic acid reported by Harvey et aP5 may be due to other roles of veratryl alcohol, such as preventing enzyme deactivation. Another possibility is that the cation radical of veratryl alcohol is stabilized at the enzyme active site, making veratryl alcohol a much more efficient mediator in the enzyme system than in model systems. After initially proposing the mediating role of veratryl alcohol, Harvey et a?1 further studied its function and showed that it accelerated the oxidation of anisyl alcohol and 4-methoxymandelic acid by preventing the formation of lignin peroxidase Compound HI. Compound TU of LiP is an inactive form of the enzyme and is formed in the presence of a large excess of oxidant in absence of readily oxidizable substrates. Compound IU can also be formed through the reaction of the native enzyme with 215 superoxide, since it is formally the ferrous dioxygen adduct. Recent work by Cai and Tien 3 8 indicated that veratryl alcohol could inhibit the accumulation of Compound JU of LiP. In the presence of veratryl alcohol and oxidant, Compound HI can be reconverted to the native enzyme. As described earlier in Chapter 2, Cl l 6 TSPPFeCl also forms a stable iron(II)-dioxygen complex (corresponding to Compound HI of peroxidases) at high pH. At pH 3, under which condition most enzymic and biomimetic studies are carried out, the Compound III complex of Cli 6TSPPFeCl is not observed. The results in Table 4.3 suggest the involvement of the Compound IU form of TDCSPPFeCl at high pH. The Table 4.3 The oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA at various pH values (yield based on substrate). pH % yield of veratraldehyde TDCSPPFeCl remaining after reaction 2 50 98 % 4 11 98 % 6 6 97 % 8 4 96 % 10 10 96 % hemin catalyst was stable but veratraldehyde was produced in very low yield at pH 10. The oxidants was probably consumed by generating Compound UI from the oxoiron(IV) porphyrin and by converting the Compound HI form back to the iron(HI) porphyrin. The Compound UI form of TDCSPPFeCl seems to be not involved in the oxidation of anisyl alcohol at pH 3. Veratryl alcohol, therefore, was not able to stimulate the oxidation of anisyl alcohol. 216 From the results of Harvey et a l 3 5 3 7 , of Cai and Tien, 3 8 and those described above, it seems certain that veratryl alcohol can indeed function as a redox mediator under certain conditions. It is, however, not an efficient mediator and it can itself be irreversibly oxidized at the same time. Inhibiting the accumulation of Compound IU and thus preventing the deactivation of lignin peroxidase is certainly another very important role of veratryl alcohol. 4.3 Experimental Cyclic voltammetry (CV) was performed in a microcell using a glassy carbon working electrode, a platinum auxiliary electrode and a silver reference electrode. The electrochemical apparatus was a PAR Model 173 Potentiostat/Galvanostat, a PAR Model 175 Universal Programmer and a Tektronix 564B Oscilloscope. The controlled potential oxidation of Poly B-411 was carried put in a two compartment cell, using glassy carbon as the working electrode, a platinum plate as auxiliary electrode, and silver wire as reference electrode (all potentials are quoted according to this standard). The potentials for the oxidation of Poly B-411 in the presence of manganese complexes was controlled at 0.80 V . The formation of manganese dioxide was confirmed by the formation of a blue color upon reaction with 0.1% benzidine in 1% acetic acid. Poly B-411 was obtained from Sigma. Veratryl alcohol and anisyl alcohol were purified by vacuum distillation. m-Chloroperbenzoic acid (mCPBA) (Fisher Scientific Company, purified grade) was used without further purification. Sigma cellulose membrane, which retains 90-99% of a cytochrome c (M.W. 12,400) over a 10 hour period, was used as the dialysis bag. It was treated in boiling water for 1 hour before use. A l l other chemicals were of reagent grade. 217 The preparation of manganese(III) malonate. The electrochemical generation of Mn(HI) malonate was carried out using the same apparatus as for the controlled potential oxidation of Poly B-411. A solution containing MnS04 (lxlO"5 mole) and malonate (5xl0"3 mole) in pH 4.5 acetate buffer (10 mL) was oxidized at a potential of 0.8 V. Solid manganese(HI) acetate (lxlO"5 mole) was added to a 0.5 M malonate solution (pH 4.5, 10 mL) to a final concentration of 1 mM. The preparation of manganese(III) malate. Enough glacial acetic acid was added separately to manganese(UI) acetate (2 mmole) and malic acid (3 mmole) to make them dissolve. The malic acid solution was then added slowly to the manganese(HI) acetate solution to give a light brown gelatinous precipitate. The precipitate was filtrated, washed with glacial acetic acid and dried under vacuum at room temperature to give a light brown powder. Elemental analysis of the compound: C: 25.82%; H: 3.05%, calculated for Mn^QHAVHA C: 25.71%; H:3.21%. The oxidation of veratryl alcohol by TDCSPPFeCl and mCPBA. The reaction mixture (6 mL 0.1 M phosphate buffer) containing veratryl alcohol (30 nmole), TDCSPPFeCl (0.05 umole), mCPBA (30 ijmole), and 4'-methoxyacetophenone (30 |j.mole) was stirred in air at room temperature for 2 hours. The reaction was stopped by extraction with methylene chloride (3x10 mL). The percentage of TDCSPPFeCl remaining after reaction was estimated from UV-vis spectrum of the aqueous layer based on the Soret band absorption of TDCSPPFeCl at 424 nm. The methylene chloride extract was washed with saturated sodium bicarbonate (10 mL), dried, and the residue acetylated with excess 1:1 acetic anhy<Wde/pyridine for 24 hours. Saturated sodium bicarbonate (10 mL) was then added and the mixture stirred for 20 minutes. Ethyl acetate (20 mL) was used to extract the organic compounds. The ethyl acetate layer was then washed in sequence with saturated sodium bicarbonate (10 mL), water 218 (10 mL), and 10% HCl (2x10 mL). The ethyl acetate layer was then dried over sodium sulphate, evaporated, and analyzed by gas chromatography. The oxidation of anisyl alcohol in the presence of varying amounts of veratryl alcohol or 1,4-dimethoxybenzene. Anisyl alcohol (30 umole) was oxidized by TDCSPPFeCl (0.05 umole), and mCPBA (30 umole) in 6 mL of pH 3 phosphate buffer in the presence of various amounts of veratryl alcohol or 1,4-dimethoxybenzene as specified in Figure 4.8 and Figure 4.9. The reaction was stopped after two hours by extraction with methylene chloride. The percentage of the TDCSPPFeCl catalyst remaining after reaction was estimated from UV-vis spectra of the aqueous layer based on the Soret band absorption of TDCSPPFeCl at 424 nm. The extraction, derivatization, and analysis of the products was the same as described above for the oxidation of veratryl alcohol. The yield of anisaldehyde and veratraldehyde was calculated from GC integration using 4'-methoxyacetophenone as an internal standard. The electrochemically mediated decolorization of Poly B-411 in the presence of veratryl alcohol. Poly B-411 (20 mL 0.005% solution in pH 3 buffer) solution containing various amounts of veratryl alcohol (see Figure 4.10) was oxidized at 0.9 V. The decolorization of Poly B-411 was followed by monitoring the absorbance of the dye at 596 nm. The formation of veratraldehyde was followed at 310 nm by UV-vis spectroscopy. The electrochemical oxidation of Poly B-411 separated from the electrode. Poly B-411 was deposited onto a piece of filter paper by dipping it into a Poly B-411 aqueous solution (0.1%). The filter paper was then air dried and attached direcdy to the glassy carbon anode. The anolyte (20 mL) contained 10 mM veratryl alcohol and 0.1 M tetrabutylammonium perchlorate (TBAP) in methylene chloride. To prevent the rapid oxidation and oxidative polymerization of veratryl alcohol, the oxidation potential 219 was set at 1.2 V (0.3 V lower than the first oxidation peak in C V of veratryl alcohol in methylene chloride) and parts of the anode not covered by the filter paper were covered with Teflon tape. 220 References for Chapter 4 1. M . Tien, and T.K. Kirk, Science, 221, 661, 1983. 2. J.K. Glenn, M . A . Morgan, M . B . Mayfield, M . Kuwahara, and M . H . Gold, Biochem. Biophys. Res. Commun. 114, 1077, 1983. 3. T.K. Kirk, and R.L. Farrell, Ann. Rev. in Microbiol. 41, 465, 1987. 4. M . Tien, CRC Critical Rev. in Microbiol. 15, 141, 1987. 5. T.K. Kirk, Phil. Trans. R. Soc. Lond. A321, 403, 1987. 6. J.A. Buswell, and E. Odier, CRC Critical Rev. in Biotechnol. 6, 1, 1987. 7. M . Kuwahara, J.K. Glenn, M . A . Morgan, and M . H . Gold, FEBS Letters, 169, 247, 1984. 8. J.K. Glenn, and M . H . Gold, Arch. Biochem. Biophys. 242, 329, 1985. 9. A . Paszczunski, Van-Ba Huynh, and R. Crawford, Arch. Biochem. Biophys. 244, 750, 1986. 10. M . Wariishi, L . Akileswaran, and M . H . Gold, Biochem. 27, 5365, 1988. 11. Y . Mino, H . Wariishi, N.J. Blackburn, T .M. Loehr, and M . H . Gold, J. Biol. Chem. 263, 7029, 1988. 12. S.D. Haemmerli, M.S.A. Leisola, and A. Fiechter, FEMS Microb. Letters, 35, 33, 1986. 13. H.W. Kern, and T.K. Kirk, Appl. Environ. Microbiol. 53, 2242, 1987. 14. P.J. Kersten, M . Tien, B. Kalyanaraman, and T.K. Kirk, J. Biol . Chem. 260, 2609, 1985. 221 15. J.K. Glenn, L . Akileswaran, and M . H . Gold, Arch. Biochem. Biophys. 251, 688, 1986. 16. F. Sundholm, and G. Sundholm, Holzforschung, 36, 71, 1982. 17. D. Limosin, G. Pierre, and G. Cauquis, Holzforschung, 39, 91, 1985. 18. A.T. Kuhn, J. Chem. Soc. Faraday Trans. 79, 417, 1983. 19. M.E . Bodini, L . A . Willis, T.L. Riechel, and D.T. Sawyer, Inorg. Chem. 15, 1538, 1976. 20. J.K. Glenn, and M . H . Gold, Appl. Environ. Microbiol. 45, 1741, 1983. 21. D.T. Sawyer, and M.E. Bodini, J. Am. Chem. Soc. 97, 6588, 1975. 22. E. Schlertzer-Rust (Ed.), Gmelin Handbuch der Anorganischen Chemie. Koordinationserbindungenz. Springer-verlag, 1980. 23. K . D . Singh, S.C. Jain, T.D. Sakore, and A . B . Biswas, Zeitschr. Kristallograph. 141, 473, 1975. 24. T. Lis, Acta Cryst. B38, 937, 1982. 25. M . H . Gold et al, unpublished observations. 26. K . Lundquist, and T.K. Kirk, Phytochem. 17, 1676, 1978. 27. M . Shimada, F. Natatsubo, T.K. Kirk, and T. Higuchi, Arch. Microbiol. 129, 321, 1981. 28. B.D. Faison, T.K. Kirk, and R.L. Farrell, Appl. Environ. Microbiol. 52, 251, 1986. 29. M.S.A. Leilosa, D.C. Ulmer, R. Waldner, and A. Fiechter, J. Biotechnol. 1, 331, 1984. 30. D.C. Ulmer, M.S.A. Leisola, and A . Fiechter, J. Biotechnol. 1, 13, 1984. 31. S.D. Haemmerli, M.S.A. Leisola, D. Sanglard, and A . Fiechter, J. Biol. Chem. 261, 6900, 1986. 32. T. Tonon, and E. Odier, Appl. Environ. Microbiol. 54, 466, 1988. 222 33. P.J. Harvey, H.E. Schoemaker, and J .M. Palmer, Collq. INRA 40 (Lignin Enzymic Microb. Degrad.), 145, 1987. 34. H . Wariishi, and M . H . Gold, FEBS Letters, 243, 165, 1989. 35. P.J. Harvey, H.E. Schoemaker, and J .M. Palmer, FEBS Letters, 195, 242, 1986. 36. F. Cui, unpublished results, 1986. 37. P.J. Harvey, H.E. Schoemaker, and J .M. Palmer, Collq. INRA 40 Qlignin enzymic Microb. Degradn.), 145, 1987. 38. D. Cai, and M . Tien, Biochem. Biophys. Res. Commun. 162, 464, 1989. 223 Chapter 5 Summary Both lignin peroxidase and the manganese-dependent peroxidase are heme proteins containing iron protoporphyrin IX as the prosthetic group. Their catalytic cycles (Scheme S.l) are typical of peroxidases whereby the native enzyme is oxidized Resting Enzyme Ligninox Lignin Compound II H 2 0 2 02-Scheme S.l Proposed catalytic cycle of lignin peroxidase by hydrogen peroxide ( a two electron oxidant) to compound I which is two oxidizing equivalents above the resting state and has been characterized as an oxoferryl irontTV) porphyrin 7C-cation radical. Like other peroxidases the lignin peroxidase and the manganese-dependent peroxidase can then bring about two subsequent one-electron oxidations arriving back at the resting state through compound II which is an oxoferryl 224 iron(TV). The complexity of the lignin degrading enzymes and of their final substrate, kgrtin, has led us to prepare simple biomimetic catalysts (A-D) which have been used to gain an understanding of the more complex natural system. A M=Fe C M=Fe B M=Mn D M=Mn The oxidation of TDCSPPMnCl (B) gives two oxidized intermediates (Scheme 5.2), possibly oxomanganese(TV) and oxomanganese(V) porphyrins. Veratryl alcohol TDCSPPMnv=0 TDCSPPMnmCl * ^ T D C S P P M n ^ O STANDING OR VA Scheme 5.2 The redox cycle of TDCSPPMnCl (VA: veratryl alcohol) 225 can reduce the oxomanganese(V) porphyrin rapidly to give the oxomanganese(IV) porphyrin and then the manganese(IH) porphyrin, mdicating that both oxidized intermediates are responsible for the oxidation of veratryl alcohol. Cl 1 6 TSPPMnCl (D) has a similar redox cycle. An additional intermediate, Cli6TSPPMn(U), was also observed. It is suggested that the manganese(U) porphyrin is formed by the reaction of the manganese(HI) porphyrin with superoxide, which is formed in the oxidation of veratryl alcohol by Cl1 6TSPPMn(HT)Cl and NaCIO. Only one oxidized intermediate was observed when TDCSPPFeCl (A) or Cl 1 6TSPPFeCl (C) was oxidized and the nature of the oxidized intermediate was not clear. The protein part of these enzymes play at least two critical roles. One is the steric protection of the high oxidation states so that the enzymes do not bring about random oxidations, particularly of the porphyrins themselves and secondly, the protein in peroxidases is known to catalyze the rate of reaction between hydrogen peroxide and the enzyme. In order to achieve steric stabilization of our biomimetic catalysts chlorine atoms have been placed on both the phenyl rings and the B-positions of the porphyrins. In addition to providing steric stabilization, which we anticipated would make our catalysts more stable, we also expected that the electron withdrawing properties of the electronegative chlorines would increase the redox potential of the metal coordinated at the center of the porphyrin which in turn should be reflected the oxidizing power of the catalysts. The interaction between the iron and manganese porphyrins and various two-electron oxidants has been studied as a function of pH. The nature of the initial ligands coordinated to the porphyrins, the ionization state of the oxidant which in turn would affect the rate of coordination to the metal center and cleavage of the oxygen-oxygen bond to generate the high oxidation state will all be dependent upon pH. In 226 addition, some oxidant-consuming side reactions such as: PorM + t-BuOOH -» PorM + CH3COCH3 + CH3OH (PorM)^ •+ H 20 2 -> PorM + H 20 + 0 2 (where PorM is a metalloporphyrin and (PorM)^^ is its two-electron oxidized intermediate) are also pH dependent and add to the complexity of the systems. Nevertheless, useful information was obtained to direct further studies for lignin model compound degradation or pulp bleaching. The stabilities of the metalloporphyrins towards excess oxidants were dramatically increased by introducing chlorines at the phenyl rings. Further chlorination of the porphyrins at their B-positions, however, did not result in additional stabilization, probably because the B-substituted porphyrins, which have much higher redox potentials, are more powerful oxidants towards both the substrate and themselves. The presence of a readily oxidizable substrate, which can rapidly transfer electrons to the highly reactive oxidized porphyrin intermediates, will decrease the rate of self-destruction. The relative stabilities of the metalloporphyrins were measured by observing the decrease in the Soret absorption (characteristic of all porphyrins) in the presence of veratryl alcohol with oxidant present in 100 fold molar excess. It was found that in the presence of veratryl alcohol, the manganese porphyrins were more stable than their iron analogues and the more highly chlorinated porphyrins (with better steric protection) were less stable than the octachlorinated porphyrins and that the metalloporphyrins were more stable at lower pH. In the presence of veratryl alcohol, the stability order of the four metalloporphyrins studied was: 227 TDCSPPFeCl > Cl16TSPPFeCl > TDCSPPMnCl > Cl16TSPPMnCl The pH dependency of the catalytic activities of the metaUoporphyrins was studied using veratryl alcohol as a substrate. The rate at which substrate is oxidized will also be dependent upon pH in a complex manner. The optimal pH at which a metaUoporphyrin/oxidant combination has its maximal catalytic activity is listed in Table 5.1 (A duplication of Table 2.1). In general, the manganese porphyrins have higher optimal pH than their iron porphyrin analogues and the hexadecachloroporphyrins (Cl16TSPPFeCl, Cl16TSPPMnCl) have higher optimal pH than their octachloroporphyrins (TDCSPPFeCl, TDCSPPMnCl). All the metaUoporphyrins have low optimal pH when using mCPBA as an oxidant. When hydrogen peroxide is the choice of oxidant, TDCSPPMnCl and Cl16TSPPMnCl are better catalysts than their iron analogues and the optimal pH for the reaction is high (pH iO). Overall the best catalytic activity was observed with TDCSPPFeCl using mCPBA at low pH. Oxidation of the oxoiron(IV) (compound U) complex of lignin peroxidase by hydrogen peroxide generates compound UI. Compound UI is formally the ferric-superoxide complex but is equally well represented by the ferrous dioxygen resonance structure: Fe(m)-CV <-> Fe(H)-02 Compound JU These complexes can also be formed by reacting the ferrous state with 02 or the ferric complex with 02. Superoxide was formed during the oxidation of veratryl alcohol and deactivated the metalloporphyrin catalysts by formation of Mn(H) or Fe(U)-02 species, 228 Table 5.1 Optimum pH for the oxidation of veratryl alcohol by various metaUoporphyrin/oxidant combinations at room temperature Metalloporphyrin Oxidant Optimal pH ai6TSPPMna mCPBA 3 a„TSFPFeCl mCPBA 2 TDCSPPMnCl mCPBA 3 TDCSPPFeCl mCPBA 2 TDCSPPMnCl NaQO 6 TDCSPPFeCl NaQO 7 a16TSPPFea t-BuOOH 5 TDCSPPFeCl t-BuOOH 6 a16TSPPMna t-BuOOH 10 TDCSPPMnCl t-BuOOH 10 al6TSPPMna NaQO 10 al6TSPPFea NaQO 10 TDCSPPFea HA 2 TDCSPPMnCl HA 10 CluTSPPMnCl HA 10 corresponding the compound IU of the peroxidases. However, addition of a superoxide dismutating agent, manganese(II)-lactate, increased the catalytic activity of the metalloporphyrin catalyst by altering the rate of compound BI formation. Addition of imidazole was also found to increase the catalytic activity of Cl16TSPPMnCl and H A for veratryl alcohol oxidation, supporting the suggestion that the ligation of the metalloporphyrin catalysts will change catalytic activity, a phenomenon which is well characterized with heme proteins. Since the electron transfer between the high molecular weight heme proteins and 229 the even higher molecular weight lignin can be expected to be very inefficient, it has been suggested that redox mediation must play an important role in the biological degradation of lignin by lignin peroxidase and the manganese-dependent peroxidase. Manganese complexes, which are essential for the activity of the manganese-dependent peroxidase, were found to function as redox mediators between the enzyme and substrates. We have used electrochemical methods to study the mediating ability of manganese complexes. The electrochemical oxidation of Poly B-411, a lignin model compound, was accelerated by addition of manganese(U)-carboxylic acid complexes. The function of the various carboxylic acids was to stabilize the manganese(JJJ) ions and to prevent their disproportion to Mn02. The optimal concentrations of lactic acid and malic acid in the electrochemical system were similar to those of the enzyme system, indicating the electrochemical system indeed mimics the enzyme system. Malonic acid was found to be the best ligand for maximal acceleration in the electrochemical oxidation of Poly B-411. We learned after finishing this work that M. Gold et al (personal communication) also found that malonic acid could dramatically stimulate the activity of the manganese-dependent peroxidase. It has been proposed that veratryl alcohol can function as a redox mediator for lignin peroxidase. It has also been proposed that veratryl alcohol can increase the activity of Ugnin peroxidase by preventing enzyme deactivation. The mediating abitity of veratryl alcohol was studied in electrochemical and biomimetic systems, where deactivation of the enzyme is not a problem and the mediating effect can be clearly studied. The electrochemical oxidation of Poly B-411 in aqueous solution was drarnaticaUy accelerated by the addition of veratryl alcohol, suggesting that veratryl alcohol was a redox mediator for the reaction. Poly B-411, when deposited on a piece of filter paper and physicaUy separated from the electrode, was also oxidized in the 230 presence of (but not in the absence) veratryl alcohol, again indicating that veratryl alcohol mediated the electrochemical oxidation of Poly B-411. Harvey et al suggested that veratryl alcohol facilitated the oxidation of anisyl alcohol and 4-methoxymandelic acid by lignin peroxidase by association of the veratryl alcohol cation radical with the compound JJ of the enzyme to make it a more powerful oxidant, therefore prevent the formation of compound III. However, in our studies the presence of veratryl alcohol was unable to increase the yield of anisyl alcohol oxidation by TDCSPPFeCl and mCPBA. While lignin peroxidase and the manganese-dependent peroxidase cannot catalyze the depolymerization of lignin polymers, they can catalyze the oxidation of a number of low molecular weight compounds representing the various substructures of lignin. We studied in detail the oxidation of a number of lignin model compounds catalyzed by one of the metalloporphyrins, TDCSPPFeCl. Veratryl alcohol, one of the simplest lignin model compounds, was oxidized by TDCSPPFeCl and /nCPBA in aqueous buffer to give products similar to that of lignin peroxidase catalyzed reactions. The oxidation of other lignin model compounds, including B-l, B-O-4, phenylpropane and phenylpropene model compounds, were also studied and the reactions were found to be similar to those catalyzed by lignin peroxidase. Almost all the reactions catalyzed by tignin peroxidase, including benzyl alcohol oxidation, benzylic methylene group hydroxylation, carbon-carbon double bond dihydroxylation, side chain carbon-carbon bond cleavage, demethoxylation, and aromatic ring cleavage reactions, were observed in the oxidation of various lignin model compounds by TDCSPPFeCl. The oxidation of model compounds representing the B-5 substructure of lignin has not been reported. TDCSPPFeCl was able to catalyze the oxidation of a B-5 model 231 compound to give products resulting from side chain carbon-carbon bond cleavage and aromatic ring oxidations. A 5-5' biphenyl model compound, representing a substructure of lignin most resistant to microbial degradation, was also oxidized by TDCSPPFeCl and t-BuOOH to give side chain oxidation and aromatic ring cleavage products. The 6-5 and 5-5* biphenyl model compounds were probably oxidized by TDCSPPFeCl through the same one-electron oxidation mechanism and the formation of the products can be rationalized by this mechanism. The isolation of novel products from the oxidation of various lignin model compounds also gives insight into the mechanism of the reactions. The formation of direct aromatic ring cleavage products from the oxidation of veratryl acetate, the phenylpropane, B-5 and 5-5' biphenyl model compounds supports the mechanism of aromatic ring cleavage previous proposed; The formation of a compound (E) similar to a proposed intermediate (F) for the phenyl-alkyl cleavage supports the proposed mechanism and indicates the necessity of the presence of a hydroxy group on the a-carbon of the side chain; The formation of the methanol incorporated products in the oxidation of veratryl alcohol and veratryl acetate in methanol suggests that the solvent molecule preferentially attacks the Cs-position of the substrate cation radical; The formation of the solvent-incorporated product in the oxidation of the phenylpropene model compound suggests a cation radical mechanism of the reaction. 0 0 E F 232 Lignin peroxidase can also oxidize environmental pollutants including polycyclic aromatic hydrocarbons and polychlorinated phenols. TDCSPPFeCl was shown able to catalyze the oxidation of pyrene and 2,4,6-trichlorophenol to give products similar to those of the lignin peroxidase catalyzed reactions. In summary, the metalloporphyrins used in this study have been shown to closely mimic the function of lignin peroxidase in the oxidation of lignin model compounds. Factors affecting the catalytic activities of the metalloporphyrins have been studied. Unlike the enzymes which are very labile, the metalloporphyrins can be used over a wide range of pH and temperature, suggesting that they may have potential industrial applications. 233 Appendix 1 The oxidation of Lignin Indulin AT Introduction Lignin Indulin AT is a commercially available industrial kraft lignin which is insoluble in water and soluble in basic aqueous solution or aqueous DMF. It is not a 'natural' lignin since the lignin structure is modified during the kraft pulping process. Initial studies of possible lignin degradation by metalloporphyrins were carried out using this easily available lignin. UV-vis spectrometry has been used in the study of lignin chemistry.1"3 Ionization difference spectra,2,3 the UV-vis spectra of a lignin sample in basic solution (when the phenolic hydroxyl groups are ionized), subtracted from that of the same sample in neutral or acidic solution, has been used to estimate the hydroxy group content of ligniris. The typical difference spectrum of softwood lignosulfonate has peaks at about 250 nm and 300 nm, respectively.3 This was explained as being due to the presence of unconjugated guaiacyl moieties (85) in lignin. The aromatic hydroxy content can be O H O H 85 86 234 estimated based on the intensity of either of these two peaks. When the hydroxyl group was conjugated with a C0-carbonyl group (86), the peak at 300 nm in the difference spectrum will shift to longer wavelength by 50-60 nm accompanied by a large increase in intensity.3 The estimation of hydroxy content by using the intensity of the long wavelength peak is no longer valid if the lignin sample contains Ca-carbonyl groups. The C0-carbonyl groups can be reduced by sodium borohydride (NaBHJ to give Ca-alcohols and aromatic hydroxy content of the sample can be determined by this so called reduced-difference spectrum. Results and discussion The oxidation of Lignin mdidin AT by a^TSPPMnCl/HjOj or Cl l 6TSPPMnCl/t-BuOOH was studied. Difference spectra and reduced-difference spectra provide a simple means to roughly predict the chemical changes after oxidation. The difference spectrum of Lignin Indulin AT has peaks at 252 nm, 300 nm, and 374 nm, respectively. The 374 nm peak was probably due to the presence of conjugated phenolic units. The difference spectra of Lignin Indulin AT treated with Cl 1 6TSPPMnCl/H 20 2 or a i 6TSPPMnCl/t-BuOOH at either pH 2 or pH 10 are distinctively different from that of the untreated sample as shown in Figures A - l to A-4, indicating extensive modification of lignin by Cl1 6TSPPMnCl catalyzed oxidation. The 374 nm peak in the difference spectra of all the oxidized lignin samples either shifted or disappeared, suggesting that the conjugated structure 8 6 was converted to other structures. In the case of lignin treated with a,6TSPPMnCl/t-BuOOH at pH 2 (Figure A- l ) , both the 300 nm and 374 nm peaks disappeared in the difference spectrum and only a negative peak was found in the reduced-difference spectrum, 235 Figure A-l Difference spectra and reduced-difference spectra of Lignin Indulin AT, — difference spectrum of untreated lignin; difference spectrum of lignin treated with Cli6TSPPMnCl and t-BuOOH at pH 2; — reduced-difference spectrum of lignin treated with Cl,6TSPPMnCl and t-BuOOH at pH 2 + 0 . 5 6 8 - 1 t o — I UJ o z < CO cr o cn CQ < + 0 . 4 3 0 -+ 0 . 2 9 3 -+ 0 . 1 5 5 -+ 0 . 0 1 8 -300 4 0 0 5 0 0 6 0 0 WAVELENGTH Figure A-2 Difference and reduced-difference spectra of Lignin Indulin AT treated with Cl16TSPPMnCl and H 20 2 at pH 2, :— difference spectrum of untreated lignin; difference spectrum of treated lignin; — reduced-difference spectrum of treated lignin Figure A-3 Difference and reduced-difference spectra of Lignin Indulin AT treated with Cl16TSPPMnCl and t-BuOOH at pH 10, ;— difference spectrum of untreated lignin; difference spectrum of treated lignin; — reduced-difference spectrum of treated lignin +0.570 +0.448-+0.326-+0.204-+0.082--0.040 700 WAVELENGTH Figure A-4 Difference and reduced-difference spectra of Lignin Indulin AT treated with G 1 6TSPPMnCl and H 2 0 2 at pH 10, ;— difference spectrum of untreated lignin; difference spectrum of treated lignin; —- reduced-difference spectrum of treated lignin indicating both the unconjugated and conjugated guaiacyl structures (85 and 86) had disappeared. Unfortunately, the oxidation of phenolic model compounds by lignin peroxidase and biornimetic catalysts have not been extensively studied. The type of structural changes that have occurred cannot be explained based on our present knowledge of these difference spectra. In addition, reactions occurred on the non-phenolic moiety of the lignin structure, such as side-chain cleavage, demethoxylation, and aromatic ring cleavage reactions cannot be seen from difference spectra or reduced-difference spectra. C 1 3 NMR, FT-IR, and molecular weight distribution analyses of the oxidized lignin sample are needed to understand any of the resulting structural changes. Experimental Kraft softwood Lignin Indulin AT, obtained from Westraw Corporation, Charleston Heights, S.C, USA, was dried under vacuum at room temperature for 24 hours before use. Lignin InduHn AT (1.0 mg) was dissolved in a mixture of DMF (1 mL) and aqueous buffer (0.1 M phosphate buffer, pH 2 or 10, 2 mL) containing Cl16TSPPMnCl (5x10-* mole). The oxidant (9 pi of 30% H A or 90% t-BuOOH) was added in three portions over 60 minutes and the reaction mixture was stirred at room temperature for an additional 24 hours. The resulting solution was diluted with water (7.0 mL) and 1 mL of the diluted solution was mixed with 0.1 M pH 6 buffer (1.0 mL) to serve as the reference for the difference spectral measurements. The sample for the difference spectral measurements was obtained by mixing the diluted solution (1.0 mL) with 0.1 M pH 12 borate buffer (1.0 mL). When the reaction was carried out at pH 2, the solution was first neutralized with 10 drops of 1 M KjCOj before diluting 240 with water (7.0 mL). For reduced-difference spectra, the diluted solution (2.0 mL) was mixed with 0.1 M pH 12 borate buffer (2 mL) and reacted with sodium borohydride (2.0 mg) at room temperature for 24 hours. The reference and scanning sample for the reduced-difference spectra was obtained by diluting the reduced solution (1 mL) with pH 6 (1 M phosphate) and pH 12 (1 M borate) buffer (1 mL), respectively. Appendix 2 Metallophthalocyanines as Possible Lignin Peroxidase Models Introduction As described in previous chapters, the chloro-substituted metalloporphyrins (TDCSPPFeCl, TDCSPPMnCl, Cl16TSPPFeCL and Cl16TSPPMnCl) are very stable and closely mimic the function of lignin peroxidase in the oxidation of lignin model compounds. Cl16TSPPFeCl has also been shown to be an effective catalyst for pulp bleaching and pulp mill effluent decolorization, and therefore has potential industrial apphcations. The metalloporphyrins are, however, expensive and cheap substitutes need to be found. The phthalocyanines (87) belong to a class of synthetic compounds closely related to the naturally occurring porphyrins4 (88). Metallophthalocyanines are known to have R 2 R« R3 87 88 242 catalytic activities for a variety of reactions5 including hydrocarbon autoxidation,6"7 olefin epoxidation8,9 and hydrocarbon hydroxylation.10 Iron phthalocyanines have also been shown to have catalase- and peroxidase-like activities.11"" The use of metallophthalocyanines as lignin peroxidase models, however, have not been previously reported. We studied the oxidation of lignin model compounds by several metallophthalocyanines, including Fe(Tfl)-, Mn(m>, Ni(U)-, Cu(U)-, and Co(H)-tetrasulfonatophthalocyanine (TSPCFeCl, 89; TSPCMnCl, 90; TSPCNi, 91; TSPCCu, 92; TSPCCo, 93; for general structure see 94). The stability and catalytic activities of these metallophthalocyanines are compared. • - SCbH Results and Discussion Veratryl alcohol (3,4-dimethoxybenzyl alcohol) is a secondary metabolite of lignin biodegradation by P. chrysosporium and is often used as a lignin model compound. The oxidation of veratryl alcohol to give veratraldehyde (3,4-dimethoxybenzaldehyde) has been used to measure the activity of lignin peroxidases and is used in this study to compare the catalytic activities of the metallophthalocyanines. 243 With TSPCCo, TSPCCu, and TSPCNi as the catalysts, the oxidation of veratryl alcohol by various oxidants gives only trace amounts of veratraldehyde as seen by thin layer chromatography (TLC). Iron(TJI) and manganese(lil) phthalocyanine are better catalysts for the oxidation of veratryl alcohol and the yields of veratraldehyde under various conditions are listed in Table 1. The yields are still too low for TSPCFeCl and TSPCMnCl to be effective catalysts. In addition, the catalysts are not stable in most cases and are bleached within a few minutes. In the oxidation of veratryl alcohol by TSPCMnCl and mCPBA, for example, TSPCMnCl is bleached within 1 minute. The catalyst can be considered to be effective since it is destroyed within one minute after the addition of oxidant and the yield is still 30% (based on veratryl alcohol), unless the degradation products of TSPCMnCl also have catalytic activity. An experiment was carried out where mCPBA was added to a solution containing TSPCMnCl, and veratryl alcohol was not added to the solution until the deep green color of TSPCMnCl disappeared. Little veratraldehyde was detected after 60 minutes, indicating that TSPCMnCl was the only catalyst and its degradation products had no catalytic activity. Table A-2. The effect of imidazole on the oxidation of veratryl alcohol by TSPCMnCl and H A at pH 7*> at room temperature irmdazole/TSPCMnCl (molar ratio) 1 10 50 100 % yield of veratraldehyde 17 20 23 10 (based on veratryl alcohol) 'h The reaction mixture contains 5x l0 7 mole of TSPCMnCl, 1x10"* mole of H 2 0 2 , and 5x10 s mole of veratryl alcohol. Peroxidases, including lignin peroxidases, have an iron protoporphyrin DC with a proximal-bound imidazole group of histidine at their active sites.1*43 Imidazole has been 244 Table A - l Yield'' for the oxidation of veratryl alcohol by TSPCMnCl and TSPCFeCl under various conditions10 t mCPBA t-BuOOH H 2 0 2 NaCIO PhIO KHS0 5 pH 3 32 3 1 3 14 0 pH 7 27 11 17 4 7 trace TSPCMnCl pH 10 29 16 10 , 6 9 2 CHjCN 24 NDC) ND ;ND ND ND CHjOH 27 ND ND ND ND ND pH3 12 3 trace 3 9 7 TSPCFeCl pH7 7 1 1 1 6 trace pH 10 16 3 1 1 4 trace yield based on veratryl alcohol, reaction carried out at room temperature b ): The reaction mixture contains in 5 mL solvent 1x10* mole of oxidant, 5x l0 7 mole of metaUophthalocyanine and 5x10 s mole of veratryl alcohol. e ): not determined used to facilitate metaUorx)rphyrin catalyzed reactions.16,17 We have shown that the presence of imidazole could increase the catalytic activity of a manganese porphyrin for the oxidation of veratryl alcohol by hydrogen peroxide. As shown in Table 2, the presence of imidazole at certain concentrations also increased the yield of veratraldehyde. The yield for the oxidation of veratryl alcohol by metalloporphyrins was dependent on the solvent The solvent effect for the oxidation of veratryl alcohol by TSPCMnCl and mCPBA was studied. TSPCMnCl was much more stable towards mCPBA in organic solvents such as acetonitrile and methanol (containing 100 uL of water in 5 mL solvent) than in aqueous solution. The yield of veratraldehyde (Table 1), however, does not change much with the solvent The oxidation of two dimeric lignin model compounds by TSPCFeCl and TSPCMnCl was also studied. The oxidation of the B-O-4 dimer, 4-ethoxy-3-methoxyphenylglycerol-B-guaiacyl ether (62), by mCPBA and either TSPCFeCl or TSPCMnCl was not complete and gave only small amounts of 4-ethoxy-3-methoxybenzaldehyde and the a-ketone 95 as shown in Figure A-5. Similar products were found in the oxidation of the B-O-4 dimer by lignin peroxidases.18 Figure A-5 The oxidation of 62 by TSPCMnCl (or TSPCFeCl) and mCPBA in aqueous acetonitrile (pH 3) at room temperature 246 The 6-1 dimeric model compound, l-(4-ethoxy-3-methoxyphenyl)-2-(4-methoxyphenyl)-l,3-propanediol (55), was oxidized more readily than the B-O-4 dimer. No starting material was detected by HPLC when it was oxidized by either TSPCFeCl or TSPCMnCl in the presence of two equivalents of mCPBA. The products were similar to those of the reaction catalyzed by lignin peroxidases and metalloporphyrins19 (Figure A-6). 5 5 9 6 Figure A-6 The oxidation of 55 by TSPCMnCl (or TSPCFeCl) and mCPBA in aqueous acetonitrile at room temperature MetaUophmalocyanines have been used as catalysts for a variety of reactions including epoxidation of olefins,8,9 hydroxylation of hydrocarbons,10 and decomposition of hydrogen peroxide.11,12 The stability of these catalysts in the presence of excess oxidants, which may be a major problem for their industrial applications, have been rarely discussed.8 Among the metallophthalocyanines used in ibis study, TSPCCu, TSPCNi, TSPCCo, TSPCFeCl, and TSPCMnCl, both of the effective catalyst TSPCFeCl and TSPCMnCl were unstable and bleached within a few minutes under the experimental conditions. One exception is that TSPCFeCl is relatively stable in the presence of hydrogen peroxide. TSPCFeCl, however, has very litde activity for the oxidation of veratryl alcohol when using hydrogen peroxide as the oxidant The stability of 247 TSPCFeCl and TSPCMnCl was dependent on the pH of the solvent as well as the type of oxidant used. As the UV-vis absorption peaks of TSPCFeCl and TSPCMnCl shifts upon addition of oxidants, it is impossible to follow the destruction of TSPCFeCl and TSPCMnCl at a single wavelength and quantitative stability data is not available. Figure A-7 shows UV-vis spectral change of TSPCMnCl and TSPCFeCl upon addition of 1 equivalent of oxidant at pH 3, 7, and 10. In the presence of a large excess of oxidant, i.e. 100 fold excess, TSPCFeCl and TSPCMnCl are destroyed very quickly. TSPCFeCl and TSPCMnCl are much less stable compared with the sterically protected metaUoporphyrins such as TDCSPPFeCl. They are also less stable than the non-sterically protected metaUoporohyrin such as me5o-terra(4-sulfonatophenyl)porphyrin iron chloride (TSPPFeCl), which retains about 30% of its activity 10 minutes after adding 100 fold excess of oxidants. TSPCCu and TSPCNi, on the other hand, are very stable and little spectral change is observed even in the presence of 100 fold excess of oxidants. However, they have little catalytic activity for the oxidation of veratryl alcohol. It can be seen by exarnining Table A-l and Figure A-7 that there is no correlation between the catalytic activity and stability of the metallophthalocyanines. Conclusions The copper(jT), nickel(IJ), cobalt(H) complexes of tetrasulfonatophthalocyanine were found to have litde activity for the oxidation of veratryl alcohol under the conditions used. TSPCFeCl and TSPCMnCl could generally mimic the function of lignin peroxidase in the oxidation of veratryl alcohol, B-l, and B-O-4 Ugnin model compounds. The presence of imidazole sUghdy increased the yield of the oxidation of 248 I Figure A-7 (C) TSPCMnCl/KHS0 3 , pH 10 + 1.341 -i +1.091 +0.841 +0.591 +0.341 +0.090 4-300 400 500 WAVELENGTH -i > ' 1 ' r 600 700 I Figure A-7 (E) TSPCMnCl/mCPBA, pH 7 Figure A-7 (F) TSPCMnCl/mCPBA, pH 10 Figure A-7 (G) TSPCMnCl/NaCIO, pH 3 Figure A-7 ( H ) TSPCMnCl/NaCIO, p H 7 Figure A-7 (I) TSPCMnCl/NaCIO, pH 10 Figure A-7 (J) TSPCMnCl/t-BuOOH, pH 3 Figure A-7 (K) TSPCMnCl/t-BuOOH, pH 7 Figure A-7 (L) TSPCMnCl/t-BuOOH, pH 10 F i g u r e A - 7 ( M ) T S P C M n C l / H 2 0 2 . p H 3 +1.370 + 1.096H +0.822 +0.54B +0.274 0.0000 300 400 500 WAVELENGTH 600 Figure A-7 (O) TSPCMnCl/H202» pH 10 Figure A-7 (P) TSPCFeCl/mCPBA, pH 3 Figure A-7 (Q) TSPCFeCl/mCPBA, pH 7 Figure A-7 (R) TSPCFeCl/mCPBA, pH 10 Figure A-7 (S) TSPCFeCl/H202, pH 3 + 0 . 6 6 4 - . o-, + 0 . 5 3 1 4 + 0 . 3 9 8 H LU U z < m c r o tr> m < +0 .266 + 0 . 1 3 3 0 . 0 0 0 0 -V 300 400 500 WAVELENGTH 600 700 800 Figure A-7 (T) TSPCFeCl/H 2 0 2 , pH 7 +0.700-1 Figure A-7 (U) TSPCFeCl/H202, pH 10 +0.560 - t=120 +0.420-ON W O 2 < m cr o tn ca < +0.2B0-+0.140 0 .0000 - r 300 400 500 WAVELENGTH 600 700 800 Figure A-7 UV-vis spectral changes of TSPCMnCl and TSPCFeCl (lxlO"7 mole in 3 mL 0.1 M phosphate buffer) upon addition of one equivalent of oxidant (lxlO"7 mole) in the presence of veratryl alcohol (5x10* mole) at room temperature over a time period of 2 minutes, 10 seconds between each scan veratryl alcohol by TSPCMnCl and hydrogen peroxide. A serious problem with these two catalysts was their low stability, which was dependent on the pH of the solvent and the type of oxidant used. The use of organic solvents instead of aqueous solvents increased their stability but did not change their overall catalytic activity. The structure of the metallophthalocyanines has to be modified to increase their stability before they can be used as effective biomimetic catalysts. Experimental Chemicals. . Veratryl alcohol, 4'-methoxyacetophenone, hydrogen peroxide, t-butylhydroperoxide, sodium hypochlorite, m-chloroperbenzoic acid (mCPBA), and potassium hydrogen monopersulfate (OXONE®) were all of reagent grade. Copper phthalocyanme-3,4^ 4/ 4^''/-tetrasulfonic acid tetrasodium salt (TSPCCu) and nickel phthalocyaninetetrasulfonic acid tetrasodium salt (TSPCNi) were purchased from Aldrich. Phmdocyaninetetrasulfonic acid was purchased from Porphyrin Products. Iodosylbenzene was synthesized as reported.20 4-Ethoxy-3-methoxyphenylglycerol-B-- guaiacyl ether (B-O-4 dimer), and l-(4-ethoxy-3-methoxy)-2(-4-methoxyphenyl)propane-1,3-diol (B-l dimer) were synthesized as described in Chapter 3. All UV-vis spectra were measured on a HP8452A diode array spectrometer. The HPLC was a Waters 600E pump with a Waters 994 photodiode array detector equipped with a 3.9 mm x 15 cm C-18 column. The mobile phase used for all analyses was 1:1 methanol/water. The synthesis of cobalt tetrasulfonatophthalocyanine (TSPCCo). Phthdocyaninetetrasulfonic acid (0.5 g) was dissolved in DMF (30 mL). Cobalt(IT) chloride (0.5 g) was added to the solution and stirred at 160 °C. The reaction was 270 complete in 60 minutes as followed by UV-vis spectroscopy. The reaction mixture was then cooled to room temperature and the product precipitated with 200 mL of acetone. The precipitate was collected, washed with acetone, dissolved in water, and filtered. The filtrate was passed through an ion exchange column (3 x 15 cm glass column packed with Amberlite IR-120 ion exchanger, H + type) to remove free Co2*. The aqueous solution was dried on rotavapor to give cobalt tetrasulfonatophthalocyanine as black crystalline powder. The UV-vis spectrum (in water) of the TSPCCo so obtained showed that it was a mixture of oxygen free and oxygen-bound cobalt complex.21,22 The synthesis of tetrasulfonatophthalocyanine manganese chloride (TSPCMnCl). Phtiialocyanmetetrasulfonic acid (0.5 ) and Mn(OAC)24H20 (1.7 g) were dissolved in water (30 mL) and refluxed for 60 minutes. A black solid was precipitated when the reaction mixture was allowed to cool down. Acetone (200 mL) was added to further precipitate the product. The precipitate was collected and washed with acetone. The manganese tetrasufonatophthalocyanine so obtained was probably a polymer and had very low solubility in water. It was dissolved in dilute sodium hydroxide, filtered, and carefully neutralized to neutral pH with dilute hydrochloric acid. The solution was then passed through ion exchange column and dried as described above. The tetrasufonatophthalocyanine manganese chloride is a black crystalline powder and dissolves in water to give a green solution. Its UV-vis spectrum is similar to that of manganese(in) phthalocyaninetetrasulfonic acid.21,23 Tetrasulfonatophthalocyanine iron chloride (TSPCFeCl) was synthesized following the method described above for the synthesis of cobalt tetrasulfonatophthalocyanine except FeClI-4H20 (0.25 g) was used. It has the same UV-vis spectrum as that reported for Iron(IU) phthalocyarimetetrasulfonic acid.21^4 The oxidation of veratryl alcohol. The reaction was started by adding the oxidant 271 (1 x 10^  moles) to 5 mL of aqueous phosphate buffer (0.1 M) contairdng 5 x 107 moles of metaUophmalocyanine and 5 x 10~5 moles of veratryl alcohol. The mixture was stirred at room temperature for 60 minutes and any remaining oxidant was decomposed by adding solid sodium bisulfite. When the reaction was carried out at pH 3, saturated sodium bicarbonate was added to neutralize the solution before adding sodium bisulfite. Saturated sodium bicarbonate was also added to the solution when mCPBA was the oxidant to remove the m-chlorobenzoic acid. 5 x 10"5 moles of 4'-methoxyacetophenone was added (as an internal standard for HPLC analysis) and the reaction mixture was men extracted with ethyl acetate (3x3 mL). The extracts were combined and analyzed by HPLC for the formation of veratraldehyde. The oxidation of fi-04 and fi-1 dimers. 5 x 10"6 moles of either B-O-4 or B-l dimer was dissolved in 4 mL 1:1 acetonitrile/pH 3 aqueous buffer containing 5 x 10"7 moles of either iron(TJI) or manganese (III) tetrasutfonatophthalocyanine, 1 x 10s moles of mCPBA was added as a powder and the mixture was stirred at room temperature for 60 minutes. Saturated sodium bicarbonate was added to adjust the pH to neutral and solid sodium bisulfite was added to decompose remaining mCPBA. The solution was extracted with ethyl acetate (3x3 mL), the organic layers were combined, evaporated to 2 mL and subject to HPLC analysis. The products were identified by comparing their retention times on HPLC, their UV-vis spectrum index, and their R, values on TLC with those of known standards. Compounds 95 and 96 were synthesized by oxidizing compounds 62 and 55, respectively, by 2,3-(licWoro-5,6^ <licyano-l,4-benzoquinone in dioxane at room temperature25. 272 References for Appendices 1. O. Goldschmid, J. Am. Chem Soc. 75, 3780, 1953. 2. O. Goldschmid, Ana, Chem. 26, 1421, 1954. 3. A.S. Wexler, Ana. Chem. 36, 213, 1964. 4. B.D. Berezin, Coordination Compounds of Porphyrins and Phthalocyanines, John Wiley & Sons, 1981. 5. FJL Moser, and AX. Thomas, The Phthalocyanines, Vol. I, CRC Press, 1983. 6. H. Kropf, Angew. Chem. intemat Edit. 11, 239, 1972. 7. CA. Tolman, J.D. Druliner, P.J. Krusic, MJ. Nappa, W.C. Seidel, I.D. Williams, and S.D. Ittel, J. MoL Cat 48, 129, 1988. 8. R. Belal, and B. Meunier, J. Mol. Cat 44, 187, 1988. 9. A.L. Stautzenberger, U.S. patent 3,931,249, 1976. 10. S.N. Massie, U.S. patent 3,692,842, 1972. 11. H. Shirai, S. Higaki, K. Hanabusa, and N. Hojo, J. Poly. Sci. Poly. Letters Edit 21, 157, 1983. 12. H. Shirai, A. Maruyama, J. Takano, K. Kobayashi, and N. Hojo, Makromol. Chem. 181, 565, 1980. 13. H. Shirai, A. Maruyama, M. Konishi, and N. Hojo, Makromol. Chem. 181, 1003, 1980. 14. L.A. Anderson, V. Renganathan, A.A. Chiu, T.M. Lohr, and M.H, Gold, J. Biol. Chem 260, 6080, 1985. 273 15. Y. Mino, H. Wariishi, NJ. Blackburn, T.M. Lohr, and M.H. Gold, J. Biol. Chem. 263, 7029, 1988. 16. J.-P. Renaud, P. Battioni, JJ7. Bartoli, and D. Mansuy, J. Chem. Soc., Chem. Commun. 888, 1985. 17. D. Mansuy, P. Battioni, and J.-P. Renaud, J. Chem. Soc., Chem. Commun. 1255, 1984. 18. T. Umezawa, M. Shimada, T. Higuchi, and K. Kusai, FEBS Letters, 205, 287, 1986 19. M Shimada, T. Habe, T. Umezawa, and T. Higuchi, Biochem. Biophys. Res. Commun. 122, 1247, 1984. 20. Organic Syntheses, Vol. 43, B.C. McKusick Editor, John Wiley & Sons, 1963, p60. 21. J. Veprek-Siska, E. Schwertnerova, and D. M. Wagnerova, Qiimia, 26, 75, 1972. 22. L.C. Gruen, and RJ. Blagrove, Aust J. Chem. 26, 319, 1973. 23. K. Fenkart, and C J H . Brubaker, Jr., J. Inorg. Nucl. Chem. 30, 3245, 1968. 24. H. Sigel, P. Waldmeier, and B. Prijs, Inorg. Nucl. Chem. Letters, 7, 161, 1971. 25. H. Decker, and E. Adler, Acta Chem. Scand. 15, 218, 1961. 

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
http://iiif.library.ubc.ca/presentation/dsp.831.1-0059840/manifest

Comment

Related Items