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Microbial degradation and determination of polycyclic aromatic hydrocarbons Li, Xing-fang 1994

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Microbial Degradation and Determination ofPolycyclic Aromatic HydrocarbonsbyXing-fang LiB.Sc., Hangzhou University, Hangzhou, China, 1983M.Sc., Institute for Environmental Chemistry, Academia Sinica, China, 1986M.Sc., Brock University, Ontario, Canada, 1989A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIES(Department of Chemistry)We accept this thesis as conforming to the required standardTHE UNIVERSITY OF BRiTISH COLUMBIASEPTEMBER 1994© Xing-fang Li, 1994In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)__________________Department ofThe University of British ColumbiaVancouver, CanadaDate -3 ‘53’DE-6 (2188)11AbstractMany polycycic aromatic hydrocarbons (PAHs) are carcinogenic, and it isimportant to understand their fate in the environment. In this thesis, the microbialdegradation of PAHs has been studied, with emphasis on the degradation of phenanthreneand pyrene. Several cultures were enriched from sea water and sediment samples obtainedfrom Kitimat Arm, B. C., an estuary environment that is contaminated by PAHs. It wasdemonstrated that microorganisms in the cultures are able to degrade phenanthrene veryquickly and to utilize phenanthrene as the sole carbon and energy source. For example,complete degradation of 100 mg of phenanthrene in 10 ml of medium is achieved 23 hrafter incubating phenanthrene with 0.5 ml of the culture and 10 ml of mineral salt mediumat ambient temperature. A strain (LY) that is capable of degrading phenanthrene wasisolated from a sea water culture and has been partially characterized.Another sixteen mixed cultures prepared from Kitimat samples readily degradedpyrene, in spite of the common perception that the microbial degradation of pyrene is verydifficult. Microorganisms in the cultures grew on pyrene, without any additional carbonsource. This is one of the few reports of microorganisms that can use pyrene as solecarbon source. The possibility of complete break-down of pyrene was studied byquantitatively measuring the amount of 14C02 produced from the mineralization of 14C-labeled pyrene. Up to an estimated 45% of the 14C-labeled pyrene that was initially addedinto the cultures was found to be mineralized to produce 14C02and H20, after 12 days ofincubation. The rest of the 14C activity is primarily accounted for metabolites. A typicalmetabolite, cis-4,5-dthydroxy-4,5-dihydropyrene, was identified in the cultures, by usingvarious chromatographic and spectrometric techniques developed during this study. Thismetabolite was present in all the cultures catalyzing pyrene degradation and disappearedonly after pyrene was completely degraded. Therefore, there was the possibility that cis4,5-dihydroxy-4,5-dihydropyrene could be used as an indicator for the in situ microbial111degradation of pyrene in the Kitimat Arm environment: cis-4,5-dihydroxy-4,5-dihydropyrene could be present because pyrene was continuously introduced to theKitimat environment and is constantly available to the microorganisms. Analysis ofenvironmental samples by using various analytical techniques showed that cis-4,5-dihydroxy-4,5-dihydropyrene is indeed present in some near shore sediment and porewater samples, in which a high concentration of pyrene is present. For the first time, thesestudies showed strong evidence for the in situ degradation of pyrene in the naturalenvironment. Radioactive tracer studies using 14C-labeled pyrene also showed thatuntreated sediment samples collected from Kitimat Arm can mineralize up to an estimated28% of pyrene.Two mixed cultures (Pysed-l and Pysed-2) were also tested for the ability todegrade benzo(a)pyrene and a standard mixture of sixteen PAHs. Some important factorsinfluencing the degradation of these PAHs were studied, including temperature, thecomposition of culture medium, the concentration of the PAHs, and the population of themicroorganisms. The degradation of pyrene at a temperature as low as 2 °C was observed.The addition of peptone and yeast extract was found to enhance the degradation of PAHs.Two methods were developed for the preconcentration of trace amounts oforganics in water samples, one based on the use of C18 solid phase cartridges the other onpolytetrafluoroethylene tubing.ivTable of ContentsAbstract iiTable of Contents ivList of Tables ixList of Figures xiList of Abbreviations xvAcknowledgments xviDedication xviiiChapter 1. Introduction 11.1. General 11.2. Toxicity 31.3. PAHs in the Environment 41.3.1. Sources 41.3.2. PAHs in water 51.3.3. PAHs in sediments 61.3.4. PAHsinbiota 81.3.5. PAH cycle in aquatic environment 91.4. Microbial Degradation of PAHs 101.4.1. General 101.4.2. Mechanisms for the microbial degradation of PAHs 101.4.3. Factors affecting the degradation of PAHs 141.5. Determination of PAHs in the Environment 161.5.1. Determination of PAHs by using capillary gas chromatography 161.5.2. Determination of PAHs by using reversed phase HPLC 171.6. Kitimat Arm Environment 18V1.7. Objectives and Overview of the Thesis 21Chapter 2. Degradation of Phenanthrene by MicroorganismsIsolated from Marine Sediment and Sea water 242.1. Introduction 242.2. Experimental 252.2.1. Instrument and analysis 252.2.2. Materials 262.2.3. Cultures 272.2.4. Degradation experiment 282.3. Results and Discussion 292.3.1. Degradation of phenanthrene by microorganismsisolated from sea water 292.3.2. Degradation of phenanthrene by microorganismsisolated from sediment 322.3.3. Isolation and the Biolog tests of strains 332.3.4. Factors influencing phenanthrene degradationby a sea water culture 34Chapter 3. Microbial Degradation of Pyreneand Characterization of Metabolites 403.1. Introduction 403.2. Experimental 413.2.1. Instrumentation 413.2.2. Chemicals and culture media 413.2.3. Cultures 423.2.4. Preservation of cultures 44vi3.2.5. Isolation and the Biolog tests of strains 453.2.6. Measurement of culture growth 473.2.7. Degradation of pyrene 493.2.8. Mineralization of pyrene 493.2.9. Acetylation and GC/MS analysis 533.3. Results and Discussion 543.3.1. Degradation of pyrene by enriched cultures 543.3.2. Mineralization of pyrene 623.3.3. Identification of metabolites 663.3.4. Degradation of pyrene by original cultures 843.3.5. Degradation of pyrene by isolated strains 883.3.6. Degradation of pyrene by preserved cultures 90Chapter 4. Mineralization of Pyrene by IndigenousMicroorganisms and Identification of aMetabolite in Sediment Samples 934.1. Introduction 934.2. Experimental 954.2.1. Sampling locations and characteristics of sediment samples 954.2.2. Mineralization of pyrene by indigenous microorganismsin sediments 984.2.3. Determination of cis-4,5-dihydroxy-4,5-dihydropyrene 994.. 3. Results and Discussion 1014.3.1. Mineralization of pyrene by indigenous microorganismsin sediments 1014.3.2. Identification and determination ofcis-4,5-diliydroxy-4,5-dthydropyrene, a metabolite fromviipyrene degradation, in the natural environment 1054.3.3. Effect of temperature 1134.3.4. Effect of culture medium and tolerance to salt 1154.3.5. Effect of pyrene concentration and microorganism population 1174.3.6. Degradation of pyrene in the presence ofcis-4,5-dthydroxy-4,5-dihydropyrene or pyrenol 121Chapter 5. Degradation and Determinationof Benz(a)pyrene and the Mixed PAHs 1265.1. Introduction 1265.2. Experimental 1275.2.1. Chemicals 1275.2.2. GC/FID analysis 1275.2.3. Degradation and determination of benzo(a)pyreneandthel6PAHs 1285.2.4. Extraction and clean-up of the sediment samples 1285.3. Results and Discussion 1305.3.1. Degradation of benzo(a)pyrene by the enriched cultures 1305.3.2. Degradation of a mixture of 16 PAHs 1335.3.3. Determination of PAHs in sediments 137Chapter 6. Preconcentration and Determination ofPolycklorinated Biphenyls (PCBs) and PAHs 140Simplex Optimization of a Capillary GasChromatography Electron Capture DetectionSystem and Solid Phase Extraction for theDetermination of PCBs 140viii6.1. Introduction 1406.2. Experimental 1416.2.1. Instrument 1416.2.2. Reagents 1416.2.3. Procedures 1426.3. Results and Discussion 1436.3.1. Simplex optimization 1436.3.2. Solid phase extraction 149II. In situ Extraction/Preconcentration of PCBs and PAHsfrom Aqueous Samples by Using a PTFE Tubing 1576.4. Apparatus and Methods 1576.5. Results and Discussion 1606.5.1. Removal 1626.5.2. Recovery and eluting parameters 1646.5.3. PAHs 1676.5.5. Application: In situ preconcentration/extraction of traceorganics from sea surface microlayer and sea water 170Chapter 7. Conclusions and Future Work 172References 176ixList of TablesTable Description Page1.1. Concentration of PAFIs in some sediment samples 72.1. Degradation of phenanthrene by three cultures I, II, and III 312.2. Biolog tests of four stains isolated from a sea water culture 342.3. Effect of Fe3 and NaCl on degradation of phenanthrene 373.1. Composition of original cultures 433.2. A summary of partial characterization of 11 isolates 463.3 Degradation of pyrene by using enriched cultures 563.4 Peak area of the metabolite (3.8 mm) produced in cultures 623.5. Contents of 14C in organic and aqueous layers 633.6 Percentage of pyrene left after pyrene incubated withthe sixteen original cultures 863.7 Peak area of the metabolite producedby the sixteen original cultures 874.1. The concentrations of the 16 PAHs in the sediments 964.2. Pyrene concentration and-4CO2 produced from the mineralizationof pyrene in indigenous sediments 1024.3. Enhancement of the mineralization of pyrene by the additionof culture Pysed- 1 1054.4. Concentration of cis-4,5-dthydroxy-4,5-dihydropyrene in samples 1124.5. Percentage of pyrene left in cultures 1175.1. Percentage of benzo(a)pyrene remained in the culture (Pysed-1) after 30 and64 days of incubation relative to the initial concentration of benzo(a)pyrene 1315.2. Ratio of benzo(a)pyrene in the culture to that in the control 133x5.3. Contents of 16 PAHs in the cultures and controlsat the given incubation time 1355.4. L/B values and the relative concentrations of 16 PAHs in the culturescompared to those in the controls 1365.5. The mean concentrations of the 16 PAHs in the sediment sample C-sed 1386.1. PCB congeners used in the present studies 1426.2. Recoveries of PCBs in water samples, obtained by using Method one 1516.3. Percentage of PCBs adsorbed on the sample containers 1526.4. Recoveries of PCBs obtained by using Method two 1546.5. Recoveries of PCBs obtained by using Method three 1556.6. Recoveries of PCBs, at various concentrations spiked into water samples,obtained by using Method three 1566.7. Effect of the length of PTFE tubing on the removal and recoveryof 10 PCBs from water sample 1636.8. Percent of PCBs removed from water sample after the samplepassed through the tubing at different flow rates 1646.9. Recovery of PCBs when different volumes of toluene was used toelute PCBs from the 8 m tubing 1656. 10. Recovery of PCBs obtained at different flow rates of the eluting solvent,10 ml of toluene 1666.11. Removal and recovery of PCBs from sea water samples that are spikedwith PCB standards 169xiLists of FiguresFigure Description Page1.1. Structure and aqueous solubility of sixteen PAils 21.2. Monooxygenase and dioxygenase mediated degradation pathwaysof PAHs by microorganisms 111.3. Pathways showing aromatic ring cleavage 121.4. Partial degradation pathway of pyrene by Mycobacterium sp. PYR-1 131.5. Maps showing Kitimat Arm, Mean aluminum smelter,and the major sampling location 202.1. Absorption spectra of the sea water culture: 10 ml of medium Ainoculated with 1 ml of culture I 302.2. Effect of amount of bacteria on the degradation of phenanthrene 382.3. Effect of culture medium on the degradation of phenanthrene 393.1. Growth curves of cultures as measured by the optical density of the culture 483.2. Schematic of apparatus for trapping 14C02 513.3. Absorption spectra of a culture (Pyw-2) 553.4. HPLC traces obtained from culture Phensed-1, culture Pyw-1,and control after 48 hr of incubation 583.5. Relative concentration of pyrene and a metabolite measured at severalintervals during incubation of pyrene and an enriched sediment culturePysed-2 593.6. Relative concentration of pyrene and a metabolite measured duringthe incubation of pyrene and an enriched sea water culture Pyw-2 613.7. Percentage of pyrene mineralized following the incubation of‘4C-labeled pyrene and an enriched sediment culture Pysed- 1 65xii3.8. HPLC/ fluorescence traces obtained from the culture, standardcis-4,5- dihydroxy-4,5-dihydropyrene, and 1:1 mixture of theculture and the standard 673.9. Absorption spectra of a metabolite in cultures and standardcis-4,5-dihydroxy-4,5-dihydropyrene obtained by using HPLCIUV 693.10. Fluorescence excitation and emission spectra of metabolite in cultures andstandard cis-4,5-dihydroxy-4,5-dthydropyrene obtained by usingHPLC/Fluorescence 703.11. GC/MS traces and mass spectrum of the acetylated productof cis-4,5-dihydroxy-4,5-dihydropyrene 723.12. GCIFID traces of the acetylated products of(a) standard cis-4,5-dihydroxy-4,5-dihydropyrene, (b) culture extract,and (c) mass spectrum of (b) by GC/MS 733.13. Fluorescence intensity of a culture supernatant at Excitation 238 nmand Emission 338 nm; and Excitation 252 nm and Emission 370 nm 753.14. HPLC/UV traces of Pyw-1-y at 2 days, Pyw-1-y at 6 days,and control at 6 days 773.15. HPLC traces obtained using UV detection at 254 nm and fluorescencedetection at excitation 260 nm and emission 370 nm 783.16. Fluorescence spectrum obtained from a metabolite in the culture(retention time 6.4 mm) 793.17. Comparison of UV absorption spectra obtained from pyrenol standard;cis-4,5-dihydroxy-4,5-dihydropyrene standard; and a metabolite inthe culture 803.18. HPLCIUV trace obtained from an original culture Phensed-2 823.19. Mass spectra of fraction I collected from preparative column,using chemical ionization and electron ionization source 833.20. Scanning UV absorption spectra obtained from a culturecontaining isolated strain, Pysed-1-y 893.21. HPLCIUV traces obtained from a regenerated culture Pysed-2that had been preserved for seven months 92xlii4.1. A map showing the location, of sampling sites 974.2. Chromatograms obtained from sediment sample STN2;cis-4,5-dihydroxy-4,5-dihydropyrene standard; and 1:1 mixture ofsample STN2 and the standard 1074.3. Comparison of UV absorption spectra obtained from cis-4,5-dihydroxy-4,5-dihydropyrene standard; and purified fraction from sediment sample STN2 1084.4. GC/MS traces and mass spectrum of the acetylation productof the metabolite isolated from the sediment sample STN2 1104.5. Effect of incubation temperature on the microbial degradation of pyrene 1144.6. Effect of culture medium on the degradation of pyrene 1164.7. Effect of initial pyrene concentration and the amount of culture onthe degradation of pyrene 1194.8. Amount of cis-4,5-dihydroxy-4,5-dihydropyrene in cultureat various intervals of incubation 1204.9. The amount of pyrene and cis-4,5-dihydroxy-4,5-dihydropyrene in cultures,measured at various intervals during incubation 1224.10. The amount of pyrene in cultures and in the control,measured at various intervals during incubation 1244.11. Relative concentration of pyrene and pyrenol in cultures at variousincubation times compared to those in the culture at incubation time 0 1256.1. Responses obtained from 39 optimization experiments asa function of GC initial temperature 1456.2. Responses obtained from 39 optimization experiments asa function of initial time, held at the initial temperature 1466.3. Responses obtained from 39 optimization experiments asa function of GC column head pressure 1476:4. Resolution obtained from 39 optimization experimentsplotted as a function of temperature ramping rate 1486.5. Schematic diagram of the preconcentration/extraction apparatus 159xiv6.6. GC/ECD traces from the determination of 10 PCBs in(a) extract from the water after passing through the PTFE tubing(b) toluene eluent from the PTFE tubing(c) the standard solution 1616.7. GCIFID traces from the determination of 16 PAHs in(a) the standard solution (10 ng of each PAH)(b) toluene eluent from the PTFE tubing(c) extract from the water after passing through the PTFE tubing 1686.8. GC/FID traces from the analysis of a sea surface microlayer sample(a) and a sea water sample (b) collected at the same location 171xvList of AbbreviationsBap benzo(a)pyrenecm centimeterECD electron capture detectorFED flame ionization detectorg gramGC gas chromatograph(y)HPAH high molecular weight polycycic aromatic hydrocarbonsHPLC high performance liquid chromatograph(y)hr hourkg kilogramL literLPAH low molecular weight polycydic aromatic hydrocarbonsm metermm minuteml milliliterMS mass spectrometry and/or spectrometerM.S. mineral saltng nanogram (10-9 g)rim nanometer (10 m)PAHs polycycic aromatic hydrocarbonspg picogram (10-12 g)PTFE polytetrafluoroethylenemicrogram (10-6 g)p.1 microliter (10-6 L)RSD relative standard deviationUV ultravioletxviACKNOWLEDGMENTSI would like to express sincere thanks to my supervisors Drs. W. R. Cullen and K.J. Reimer for their guidance, support, encouragement, and enthusiasm during my Ph.D.program at the UBC. I would also like to thank my guidance committee members, Drs. G.Eigendorf, J. P. Kutney, and A. P. Wade for their helpful suggestions.I would like to specially thank Mr. G. Hewitt for his patience, friendship, usefuldiscussions, and technical support. My gratitude is also extended to the people who sharethe biological service facility, particularly those in Dr. Kutney’s group, for their cooperation and friendship. I thank my colleagues in Dr. Cullen’s group for their cooperation and many fruitful discussions. I am also thankful to a summer student, MissMonica Chu, for her help during the summer of 1992.I would like to express my thanks to a few special people at the University ofAlberta: Dr. Julia Foght in the Department of Biological Sciences for her collaboration;Dr. Norm Dovichi in the Department of Chemistry for allowing me to use his facilities;and people in Dr. Dovichi’s group for providing friendly and enjoyable atmosphere duringmy thesis writing.The completion of this thesis has special meaning to me. During the early part ofmy thesis work, my daughter Connie accompanied me, shared the excitement of myresearch, and went through all the difficulties with me at the very beginning of her life. Herbraveness and strength in fighting for life give me the strongest courage to succeed in mystudy. As she is growing into a big healthy girl, she is my continuous source of strengthand joy. No word can express my thanks to my husband, Xiao-Chun. His endless love andpatience for me always make my study easier; his enthusiasm for searching newknowledge inspires me to go forward in the future. I am very grateful to my parents foralways believing in me and encouraging me to pursue a career in science. I must thank myparents-in-law for their support and for looking after Connie during their visit, whichxviiallows me to work in the laboratory for many extra hours without worry. Special thanksgo to my brothers and sisters and friends Sharon, Suzanne, and Bob for their constantencouragement, love, and support in many ways.Finally, my thanks go to the Department of Chemistry, the University of BritishColumbia for providing me the opportunity to study, and to NSERC for the post-graduatefellowships.xviiiDEDICATIONTo my grandmother, on her 90th birthday1Chapter 1. Introduction1.1. GeneralPolycycic aromatic hydrocarbons (PAHs) consist of two or more fused aromaticrings in linear, angular, and/or cluster arrangements (1). Heterocyclic aromatichydrocarbons result from the substitution of carbon atom in the benzene ring with otherelements such as nitrogen, oxygen, and sulfur. PAHs and heterocyclic aromatichydrocarbons along with ailcylated PAHs are recognized as priority pollutants. The sixteenPAHs listed by the Environmental Protection Agency (EPA) of the United States (2) arecommonly studied for environmental assessment. Some of the parameters of these sixteenPAHs including structure and solubility in water, are shown in Figure 1.1. Many PAHs arealso listed as national priority toxic substances in the Canadian Environmental ProtectionAct (CEPA) (3), and as hazardous substances listed by the World Health Organization(WHO) (4). The WHO Expert Committee on the prevention of cancer reported in 1964that PAHs in water supplies were potential hazardous carcinogens (5). Studies on theimpact of PAHs on the environment have attracted much attention, because there is alarge number of this class of compounds present in the environment (1, 6-8).The most distinct characteristic of PAHs is their aromaticity, and the extended telectron system results in chemical stability, which is why they tend to retain the ivelectron conjugated ring systems in reactions. The it-electron systems also give distinctspectroscopic properties to PAHs which are used in the analytical measurement of PAHsin environmental samples (4, 9, 10). Hydrophobicity is another important property ofPAHs, and because of their low solubility in aqueous environment, PAHs tend to associatewith particles and eventually sink into soil and sediment.2(1) Naphthalefle(2,000-31,700)(2) Acenaphthylene—I(3) Acenaphthefle(3,900)OThC(4) Fluorene(1, 685-1, 980)(5) Phenanthrene(1, 002-1,290)(6) Anthracene(30-73)(7) Fluoranthefle(206-260)(8) Pyrene(132-17 1)n(9) Benz(a)aflth1aCe1(9.4-14)(10) Chrysene(1.8-2)rfl(13) Benzo(a)pYrefle(3-3.8)(15) Benzo(g,h,i)perYlefle(0.26)(16) Ideno( 1 ,2,3-cd)pyrene(12) Benzo(k)flUOraflthefle(14) Dibenz(a,h)aflthraCefle(11) Benzo(b)fluorantheneFigure 1.1. Structure and aqueous solubility (nglml) of sixteen PAHs31.2. ToxicityInterest in the toxicity of aromatic hydrocarbons can be dated back to 1761, whenDr. John Hill, an English physician suggested that smoking may be a possible cause ofhuman diseases (11). Chemical carcinogenesis associated with PAHs was first reported bySir P. Pitt, who studied scrotal skin cancer in English chimney sweepers (12). The harmfuleffects of soot, tar, and pitch attracted great attention, leading to the identification ofPAHs in pitches as carcinogens in the 1930’s (13). Since then the general awareness of thetoxicity of PAHs has resulted in a number of studies. By 1976, more than 30 PAHs andseveral hundred PAH derivatives were reported to cause carcinogenic effects (14). PAHsare now known to be the largest group of chemical carcinogens (15). Among the sixteenPAHs that are designated as priority pollutants by the US EPA (Figure 1), eight areconsidered to be possible carcinogens. They are benz(a)anthracene, chrysene,benzo(b)fluoranthene, benzo(k)fluoranthene, benzo(a)pyrene, indeno(l ,2,3,c-d)pyrene,dibenzo(a,h)anthracene and benzo(g,h,j)perylene (16).The effects on humans of carcinogenic PAHs in the environment has recently beenreviewed (17). The potential exposure to carcinogenic PAHs is usually estimated byexamining the concentrations of the particular PAHs in those parts of the environmentclosely associated with human activities. Chronic toxicity is of the most concern. There isa 20 to 30 year latent period between initial exposure to a carcinogen and the appearanceof resulting human cancer. In order to assess the potential carcinogenic effect of achemical without waiting for 20 years for the appearance of human cancer, the Ames test,based on bacterial reverse mutation in Salmonella, is often used to examine themutagenesity of a chemical (16, 18-20).The mechanisms on the action of chemical carcinogens are not well understood.The association of carcinogens with important biological molecules such as DNA andRNA has been proposed (21-25). Animal studies have demonstrated that PAFIs are4metabolized by liver mixed function oxidases (monooxygenase) to epoxides, dihydrodiols,phenols and quinones. These metabolites have been identified as mutagenic andcarcinogenic agents. In particular, electrophilic metabolites such as arene oxides, havebeen found to bind to DNA (25-29). Therefore, the alteration of the structure of DNAthrough the reactions with oxygenated metabolites of PAHs are likely to cause mutationsand to damage biological functions. Reactions of PAHs with proteins may also beresponsible for the carcinogenic effect of some PAHs (22, 30).Information on the acute toxicity of PAHs is very limited. Generally, the acutetoxicity increases as the molecular weight of the PAHs increases. For example, the acutetoxicity of 1-, 2-, and 3-ring aromatic hydrocarbons to fish increases by 10 fold when themolecular weight increases by 40 to 50 units (5). The oral LD50 for rat or mouse is 1780mg/kg for naphthalene, 700 mg/kg for phenanthrene, and 50 mg/kg for benzo(a)pyrene(18, 19).1.3. PAHs in the environment1.3.1. SourcesIt is generally accepted that the most PAHs in the environment are produced bypyrolysis or incomplete combustion of organic materials (3 1-34). The specific mixture ofPAHs formed depends on the temperature of pyrolysis. Unsubstituted PAHs are the mainproducts when the pyrolysis temperature is higher than 2000 °C. At intermediatetemperatures, for example 400 to 800 °C, alkylated PAHs are produced but withunsubstituted PAHs as the major components. In contrast, alkyl PAHs with two or threearomatic rings are the major components produced at low temperatures, typically 80 to150 °C (35), and they are also the major components of aromatic compounds in crudeoil..5PAHs present in the environment are derived from both natural sources andanthropogenic sources. Natural sources include volcanic eruptions, forest and prairie fires.Biosynthesis of PAHs by algae, bacteria, and plants were observed in laboratory studies(5). However, the possibility of biosynthesis of PAHs in the natural environment is stilldebatable.Anthropogenic contributions are doubtlessly the most significant sources of PAHsto the environment (5, 36, 37). Fossil fuel burning for the generation of power and heat,industrial effluents and by-products, incineration of waste, combustion in automobiles, andfood cooking are examples of anthropogenic PAH sources (38). The burning of fossilfuels is the principal source (39-42). PAH concentrations in soil samples from industrialcountries, particularly in urban areas, have been found to increase over the last 100-150years as a result of anthropogenic combustion activities (40-46).General aspects of the sources, distribution, fate, and biological effects of PAHs inthe aquatic environment have been discussed by Neff (1). The distribution of PAHs andtheir transport through environment has been widely studied over the last two decades,and detailed information is presented in a number of reviews (1, 5, 47-52).1.3.2. PAHs in waterThe concentration of PAHs in water bodies is generally very low because of thehydrophobicity of PAHs. Reliable analytical data on PAH concentrations in fresh watersystems are limited (5, 52), and much less is known about marine waters. Availableinformation has been compiled in a few reviews (6, 49-53), and it appears that the levels ofPAHs vary greatly depending on the water source and the condition of the environment.Generally the concentration of PAHs in various water bodies decreases in the followingorder: industrial or domestic effluents > surface water > drinking and ground water (6).For example, the concentration of benz(a)pyrene was found to be as high as 100 igfL in6industrial and domestic effluents (49, 54), 0.1 to 830 ngfL in surface water (17), and 0.1to 62 ng/L in drinking and ground water (17, 53).The solubility of PAHs decreases as the number of aromatic rings increases. Forexample, the aqueous solubility of phenanthrene, pyrene, benz(a)pyrene (Bap) is 1.29,0.14, and 0.0038 mgfL, respectively (52). As the number of aromatic rings increase from 3for phenanthrene to 5 for Bap, the aqueous solubility decreases by almost 3 orders ofmagnitude. Thus water samples generally contain higher amount of low molecular weightPAHs with 2 and 3 aromatic rings, and much less of the high molecular weight PAHs with4 or more aromatic rings (52, 55). The levels of different PAHs occurring in a water bodyalso depends on the sources of PAH contamination. In oil-spill sites, low molecular weightPAHs, specifically naphthalene, phenanthrene, and anthracene, predominate (52, 55, 56).This is because they are the principal PAHs in crude oil and refined petroleum. Thepresence of other hydrocarbons such as alkanes also increase the solubility of these PAHsin the contaminated water.High molecular weight PAHs with four or more aromatic rings in water samplesare often associated with particles and sink into sediment. The concentrations of thesePAHs in water samples are often expressed as the sum of the PAHs dissolved in water andthe PAHs adsorbed on the particles. It was reported that temporal variations in theconcentration of high molecular weight PAHs are correlated to rainfall, and surface runoff is the major source of these PAHs (57, 58).1.3.3. PAHs in sedimentsThe presence of PAHs in sediment is ubiquitous. Even the sediments from Arcticsites were found to contain PAHs (59). Johnson and Larson (60) made a worldwidecomparison of the total concentration of PAHs in marine and fresh water sediments. ThePAH levels ranged from 5 ng/g for undeveloped areas in Alaska to 1.79 x 106 ng!g for anoil refmery outfall in Southampton, England. Sediments from other industrial areas ranged7from 198 to 232,000 ng/g. The results of some other studies from several locations in theUnited States and Canada are summarized in Table 1.1. A comparison of the data in Table1.1 with the background level (5 nglg) reported by Johnson and Larson (60) indicates thatKitimat Arm is highly contaminated by PAHs, although direct comparison can not makewithout care.Table 1.1. PAH concentrations in sedimentsLocation PAH concentrationjig/g (dry sediment) PAHs studied ReferencesBoston Harbor, USA 0.48-718 [61]Eagle Harbor, Puget SoundWashington, USA 1,100-2,900 N-containing PAHs [62]Sydney HarborNova Scotia, Canada 0.0029-310 18 PAHs [50,63]Vancouver Harbor, B.C. 228 benz(a)pyrene [59]Arctic 0.02-33 benz(a)pyrene [59]Kitimat ArmBC, Canada 85-528 16 PAHs [64-66]The composition of PAHs within the sediments can be altered by degradation anddiagenic reactions (67-69). Degradation leads to the loss of some components, a topicwhich wifi be discussed later. Diagenic reactions lead to the in situ production ofparticular compounds, such as perylene (68). Hydrocarbon composition is also a functionof the particle size and type of sediments (70). Prahi and Carpenter (71) reported thatsmall size particles trapped in water column contained higher levels of PAHs than largesize particles. The concentrations of PAHs were found the highest near sediment and8water interface and decreased with depth in sediment cores. A similar phenomenon wasalso found in sediment cores obtained from Kitimat Arm (72). Tn an oil-spffl area, thecomposition of sedimentary PAHs largely depends on the type of oil and theenvironmental conditions such as weathering and biological activity. This was clearlydemonstrated in studies of a large number of sediment samples following the ExxonValdez oil spill (9, 73).Sedimentary PAHs reflect, to some extent, the levels of contamination of aquaticorganisms by PAHs. Dunn (74) studied three types of marine samples including sediments,bivalves, and seaweed, from ten locations in Vancouver Harbor, B. C., Canada. It wasfound that the levels of benz(a)pyrene in sediments generally correlated with those inmussels and seaweed. The results suggest that the levels of PAHs in marine organisms aredependent on the degree of contamination in sediments.1.3.4. PAHs in biotaThe accumulation of PAHs in biota is one of the major features of theenvironmental PAH cycle, having a direct or indirect impact on many living organisms (48,51, 56, 75-80). The levels of PAHs in field-sampled organisms vary widely with thesampling site and with species. A general picture can be obtained by considering the levelsof benz(a)pyrene as an indicator. In three widely divergent groups of organisms, bivalves(17, 50,76, 79, 80), crustaceans (17), and fishes (17, 56,77, 78, 80, 81), the typical levelsof benz(a)pyrene measured are 8.0-530, 0.15-9.0, and 10 jig/kg (dry weight), respectively.Bivalves have generally been found to contain the highest concentration of PAHs.The elimination of PAHs by organisms can take place when they are removed fromthe contamination source (81). This is because some organisms such as fish andcrustaceans contain necessary enzymes, such as cytochrome P-450s, to oxidize PAHs.Bivalves are believed to lack the enzymes to metabolize PAHs and thus they have apotential to accumulate PAHs. For this reason, mussels were chosen as an indicator in the9Global Ocean Monitoring Program (51, 79, 82), to assess the levels of PAHs in the marineenvironment.1.3.5. PAH cycle in aquatic environmentThe cycle of PAHs in the aquatic environment was described by Neff (1). PAHsenter the environment through many routes such as oil spill, run-off from roads, sewage,effluents from industrial processes, and fall-out from the atmosphere. Atmosphericdeposition is the principal route of PAHs to soils and long-range transport of PAHs toremote areas (5, 45). PAHs entering water from various sources quickly become adsorbedto organic and inorganic particulate matter. Much of the particulate PAHs is deposited inbottom sediments. Leaching or biological activity in the sediments may return a smallfraction of sedimentary PAHs to the water column. PAHs are readily accumulated byaquatic biota to levels higher than those in surrounding water. Relative concentrations ofPAHs in aquatic ecosystems are generally highest in the sediments, intermediate in aquaticbiota, and lowest in the water column.Routes for the removal of PAHs from the water column include volatilization fromthe water surface (mainly low molecular weight PAHs), photooxidation, chemicaloxidation, microbial degradation, and sedimentation. These mechanisms have been studiedin both the laboratory and field (83-86). Laboratory studies suggest that microbialdegradation and evaporation are removal mechanisms for low molecular PAHs, andsedimentation and photochemical oxidation for higher molecular PAHs (> 4 rings). Thefield studies did not completely agree with the laboratory studies. For example, UV lighthad a minor effect on PAHs in the field (86), whereas photochemical oxidation was alsoan important degradation mechanism for PAHs in surface water in the laboratory studies(83, 84). High molecular PAHs remained much longer periods of time in field water thanin the laboratory studies (83, 86). While there is no agreement on the relative effects of thevarious physical, chemical, biological processes on PAHs in aquatic environment, there is10a general consensus that the primary removal mechanism for PAHs from the water columnis sedimentation, and microbial degradation for sedimentary PAHs.1.4. Microbial degradation of PAHs1.4.1. GeneralThe microbial degradation of aromatic hydrocarbons was first described byStormer (87) in 1908: he isolated Bacillus hexacarbouorum which was capable of utilizingtoluene and xylene as the sole carbon source. In the following years, a number of bacteriacapable of growing on aromatic hydrocarbons were isolated (88-90). Gray and Thornton(90) found that bacteria with the capacity to oxidize naphthalene, phenol, and cresol werewide-spread in soils. By 1946, nearly 100 species of bacteria, yeast, and fungi representing30 genera were observed to have the ability to degrade various hydrocarbons includingaliphatic, olefmic, and aromatics (91-93). These initial observations paved the way fordetailed studies of the microbial degradation of PAils.Comprehensive information on the bacterial, fungal, and algal metabolism of lowmolecular weight PAils has been presented in a number of reviews (94-97). However, thedegradation of high molecular weight PAHs is less studied and poorly understood. For along time, PAHs with four or more aromatic rings were considered to be resistant tomicrobial degradation. It has only recently been confirmed that bacteria are able todegrade tetracyclic PAHs such as fluoranthene (98, 99a, 99b) and pyrene (100, 101). Thedegradation of high molecular weight PAHs is an important topic of study because of thecarcinogenicity of these compounds (17), and the growing interest in their bioremediation(102-104). The advances in the studies of microbial degradation of PAils are accordinglymarked by the discovery of PAHs with increasing number of aromatic rings that can bedegraded, which are reviewed in a number of literetures (94-97, 105-106).111.4.2. Mechanisms for microbial degradation of PAHsThe microbial degradation of PAHs is known to follow two typical enzymaticpathways: which involve either monooxygenase or dioxygenase mediated degradation.Monooxygenase enzymes are present in fungi, mammals, and bacteria. In the initialoxidation step involving monooxygenase, one atom of the molecular oxygen isincorporated into the aromatic compound (94, 95). Bacterial degradation of PARs, on theother hand, usually involves the use of dioxygenases: both atoms from molecular oxygenare incorporated into the aromatic compound. These pathways are illustrated in Figure1.2. Oxygen is essential for initial ring oxidation of PAHs in both mechanisms. Thefollowing discussion is focused on the bacterial degradation pathway: the dioxygenasepathway.Q-GIuc oside0-GlucuronideOH ..——pPhenol2It2O 1Th-OHEpixide lJç. OHy&oase P Htrans-Dt,y&odlolPoiycyclic NADAroinat OH i 1HydrOcarbon OH DehydrogenaseH pNADH+Hcs-Diydrodio CatecholFigure 1.2. Typical monooxygenase and dioxygenase mediated oxidation of PAHs bymicroorganismsNaphthalene dioxygenase has been identified as a multi-component enzyme systemthat Consists of a flavoprotein, a ferredoxin, and an iron sulfur protein (107-1 14). In the12initial step of bacterial degradation, incorporation of both atoms of the molecular oxygeninto the PAH by the dioxygenase leads to the formation of cis-dihydrodiol (115, 116). Inthe second step the cis-dihydrodiol is transformed to a dihydroxylated intermediate by adehydrogenase. This reaction is highly stereoselective (117), and is essential for furtherring cleavage (118, 106). Following the rearomatization, the aromatic ring is enzymaticallycleaved by dioxygenase enzymes at the positions either between or adjacent to the twohydroxyl groups (96) as shown in Figure 1.3.LLOHC a tech aI2.HydroXyrfluCOflicsemialdehydeFigure 1.3. Pathways showing aromatic ring cleavage.Pathways of bacterial degradation of low molecular weight PAHs such asnaphthalene. phenanthrene, and anthracene have been elucidated (94, 95, 105). Thecommon metabolites are polar molecules such as salicylic acid, catechol, phthalic acid, andprotocatechuic acid (94, 95, 105, 119). Naphthalene, for example, initially incorporatestwo oxygen atoms into the aromatic ring by dioxygenase to form cis-l,2-dihydrodiolnaphthalene, which is subsequently oxidized to salicylic acid and catechol. Furthermineralization is carried out to produce C02 and H20 (95, 1.05).OHc,s cisMuconi(acidHO13While degradation pathways of these low molecular weight PAHs are wellunderstood, only a few bacteria have been isolated, which are able to degrade tetracydicPAHs such as pyrene and fluoranthene (98-101). So far, very few pure cultures, forexample, Rhodococcus sp. UW1, were found to utilize pyrene as sole carbon source(100). Degradation of pyrene by Mycobacterium sp. PYR-l has been studied in detail(101). Mycaobacterium sp. PYR-1 is able to degrade pyrene only when organic nutrientsare supplemented in the medium, and cis-4,5-dihydrodiol pyrene was identified andisolated from the culture. Heitkamp et al. (101) proposed that this metabolite resultedfrom an initial ring oxidation at carbon atom 4, 5- positions of pyrene. A few other ringcleavage products were also identified. Based on these results, a partial bacterialdegradation pathway of pyrene was proposed (106), as illustrated in Figure 1.4. It is clearthat the degradation pathways of a high molecular weight PAH such as pyrene are morecomplicated than those of low molecular PARs.X616D ±FxC9X..00?0H [gJ COOH4.PhenanthroateCO2OHI C’0OHç.) O<,Kiyohara Pathway.—CO0H- [.9J... COOHPhihalate...- COOH0Cinnamate© OHOH44lydrOxyperinaphtheflOfleFigure 1.4. Partial pathways for the degradation of pyrene by Mvcobarerium sp. PYR- 1141.4.3. Factors affecting the degradation of PAHsThe microbial degradation of PAHs is greatly influenced by a number of factorssuch as the physical and chemical properties of the PAHs, microbial adaptation or preexposure of microorganisms to PAHs, and the physical conditions of the environment suchas temperature, oxygen availability, nutrients and pH. Different consortia ofmicroorganisms may contribute to the degradation of PAHs in the natural environment(120). However, it has been recently demonstrated that degradation of PAHs in coastalsediments is mainly due to bacteria, and yeast make minor contribution (121).Some studies have demonstrated the molecular weight and structural-dependenceof the microbial degradation of PAHs (122-124). As the number of aromatic ringsincreases, the degradation rate decreases. It is also reported that alkylation of PAHsreduces the degradation rate significantly (122, 123). Cemiglia et al.(105, 106) haveexamined the half-lives of PAHs of different structures. It is generally found that thepersistence of PAils can be presented in the following order: naphthalene < phenanthrene<methyl-naphthalene <pyrene <methyicholanthrene <benz(a)pyrene.It has been reported that chronic exposure of microorganisms to chemicaltoxicants results in a higher tolerance to toxicity and an enhanced ability of themicroorganisms to utilize the chemicals as substrates for metabolism or co-metabolism(125-129). This phenomenon is commonly observed in the degradation of low molecularweight PAHs such as naphthalene, phenanthrene, and anthracene in oil-contaminatednatural waters and sediments. For example, the degradation of naphthalene in petroleum-contaminated sediments (turnover time 7.1 1w) is much faster than that in a pristineenvironment (129). It has been proposed that chronic exposure to PAHs may result in (i)elevated total population of microorganisms, (ii) selective increase in the population ofmicroorganisms containing PAH-degrading enzymes, and (iii) induction of PAil-degradingenzymes in some indigenous microorganisms (105). Some report (125, 127) that thepopulation of heterotrophic microorganisms (those responsible for the degradation of15organic compounds) is correlated with PAH degradation rates. Others (126, 128) arguethat the total population of heterotrophic microorganisms is not a reliable indicator ofPAH degradation, because many heterotrophic microorganisms do not degrade PAHs.The actual mechanism for the enhancement of PAH degradation due to chronic exposureis not well understood.The temperature has a direct effect on the rate of PAH degradation bymicroorganisms in natural ecosystems (130). Bauer and Capone (127) reported that themineralization rates of anthracene at 20 °C and 30 0C were doubled and tripled,respectively, compared to 10 °C incubation. Similarly, naphthalene mineralization rates at20 °C and 30 °C were 2.7 and 4.6 fold, respectively, greater than that at 10 °C. Anumber of investigators (122, 125, 131, 132) have studied the seasonal change inmicrobial activity and PAH degradation rates in the natural samples, and they found thehighest in summer and the lowest in winter. This seasonal temperature effect may be dueto (i) seasonal selection of psychrophilic (low temperature-living, 10 OC) or mesophilic(moderate temperature living, 30 0C) hydrocarbon-degrading microorganisms, (ii) lowtemperature suppression of the degradative capacity of the PAFI-degradingmicroorganisms at stable populations (105), and (iii) direct effects of temperature on theenzymatic reactions.Oxygen is essential for the biodegradation of PAHs, as discussed above. A fasterdegradation of PAHs has been observed in the upper surface sediments than in thesubsurface layers because of the depletion of oxygen in the sediment below surface (133).Although anaerobic degradation of some single-aromatic compounds such as benzene andtoluene has been reported (134-136, 136a, 136b), similar degradation has not beenobserved for PAHs. However, it has been reported (127) that some aerobicmicroorganisms that exist under anoxic conditions may help reduce PAH toxicity inanaerobic sediments that are occasionally aerated by physical mixing or benthicphotosynthesis.16Other environmental factors such as pH and nutrients are also important to the- microbial degradation of PAHs. Proper nutrient and pH conditions are required for theoptimum growth of PAH-degrading microorganisms. Detailed information can be found ina number of reports (102, 137-143).1.5. Determination of PAHs in the environmentWidespread concern over environmental pollution by PAHs and their carcinogeniceffects has emphasized the need for measurements of their concentrations in a wide varietyof samples (2). Chromatography, mass spectrometry, and a number of spectroscopicmethods have been extensively used for this purpose (1, 4, 144).Environmental samples often contain a complicated matrix, and the compositionsof PAHs vary greatly with the sampling sites. In order to adequately assess the fate andimpact of PAHs on an environment, the concentration of individual PAHs rather than thetotal concentration of mixed PAHs should be determined, because the toxicity variesgreatly with the structure and molecular weight of the PAHs as discussed earlier.Consequently, a number of chromatographic techniques have been successfully developedto meet this requirement. The most widely used techniques are capillary gaschromatography (capifiary GC) and reversed phase high performance liquidchromatography (RP-HPLC). A brief discussion on these two techniques is given in thefollowing sections.1.5.1. Determination of PARs by using capillary gas chromatographyGas chromatography (GC) with capillary columns has become a routine analyticaltechnique. Analytical methodologies based on the use of GC have been validated by EPAfor the determination of PAHs in environmental samples (145). The wide application ofcapillary GC to the determination of PAHs is attributed to the development of a variety of17stable capillary columns with high separation efficiency. Many parameters of capillarycolumns have been studied (4, 146, 147) and these are the deactivation of the surface ofthe capillary tubing (148), film thickness of the coating (149), and the properties of thestationery phase (coating) (146). Fused silica capifiary columns that are coated withvarious stationary phases are widely used. Commercially available columns such as DB-1,SPB-1, and SE-30 are coated with methylsioxane; and DB-5 and SPB-5 are coated with94% methylsioxane, 5% phenyl and 1% vinyl methylsioxane. DB-5 type columns havebeen recommended in EPA methods (145, 150). Cross-linked silicone polymers have beenfound to be useful (151). By preparing proper cross-linked phases, the selectivity of thecolumn is improved (152). The cross-linked columns have a high thermal stability and anon-extractable column coating so that the background is reduced and a better detectionlimit can be achieved (153). Specific capillary columns suitable for the separation of PAHshave also been developed. These columns are coated with 5% diphenyl and 95% dimethylsioxane. Commercial columns such as PTE-5 from Supelco, HP5-MS from HewlettPackard, DB-5MS and DB-5.625 from J & W are some examples, and they arerecommended in EPA methods 625, 1625, and 827.Flame ionization detector (FID) and mass spectrometry (MS) are often used inGC. Fl]) has the advantage of low cost with reasonably good sensitivity. The combinationof capillary GC with MS detection (GCIMS) provides mass spectrum in addition tochromatographic retention information. Therefore, GCJMS has been widely used todetermine parent PAHs and PAH derivatives. It has also been used to study themetabolites of biodegradation of PAHs and to study the mechanisms of carcinogenesis ofPAHs by identifying DNA adducts (154, 155).1.5.2. Determination of PAHs by using reversed phase HPLCThe separation of PAHs by using Cig reversed phase high performance liquidchromatography (RP-HPLC) was first demonstrated in 1971 (156). Since then, the18number of applications of RP-HPLC to PAH analysis has been continuously increasing(157, 158). RP-HPLC with UV absorption and fluorescence detection is widely used forthe determination of parent PAHs and their metabolites in a variety of samples such aswater, sediment, and biological tissues (9, 16, 56, 78, 157, 159-162). This technique isalso recommended in EPA method 550.1 for the determination of PAHs in drinking water(163), method 610 for waste water, and method 8310 for solid waste (145). Thenumerous successful applications of RP-HPLC to the determination of PAHs have beendocumented in the 1993 CRC Handbook of Chromatography devoted to the liquidchromatography of polycyclic aromatic hydrocarbons (164).HPLC can be used for direct analysis of thermally labile and polar compounds inaqueous samples, whereas GCIMS is often not a suitable choice unless the compounds arederivatized and so become stable and volatile. The separation efficiency of HPLC can bemaximized by selecting a column consisting of proper packing materials and by using asolvent gradient program (165), thus different isomers of PAHs can be completelyseparated and determined (166-168).The absorption and fluorescence spectral characteristics of PAHs are very useful inanalysis. Fast and simple techniques based on HPLC/absorption and HPLC/fluorescencehave been developed (9, 56, 73, 78, 162) for screening aromatic compounds inenvironmental samples, such as water, sediment, and fish, collected from the areas affectedby the Exxon Valdez oil-spill in 1989.191.6. Kitimat Arm environmentThe present study area, Kitimat Arm, is located in Northern B. C., Canada, asshown in Figure 1.5. An aluminum smelter, Mean, is located at the head of Kitimat Armand west of the mouth of the Kitimat River. In the Alcan smelters, the Soderberg processis the primary source of aluminum and PAHs. The aluminum is produced by electrolysis byusing electrodes consisting of coal tar and pitch. This tar mixture contains large amountsof PAils, and the aluminum smelting process generates additional PAHs. Scrubbers areused to control atmospheric pollution from the Soderberg process, but the scrubber liquoris a source of PAils when discharged into receiving waters. Alcan has two major lagoonsto collect effluent discharge. The B lagoon collects discharge from wet scrubbers, coolingsystems and surface runoff. The effluent exits the lagoon through a weir, and flows downa ditch to the harbor. The lagoon consists of two ponds that are partially separated by asill that does not reach the surface. The D lagoon collects cooling water and surface runofffrom the coke calciner area, and is essentially a small settling pond. These two lagoons arethe important point sources of PAHs (64).Many aspects of the Kitimat Arm environment have been studied. The physical andbiological characteristics of the Kitimat estuary are reviewed by Bell and Kaliman in 1976(169). Possible effects due to industrial developments, including the Alcan smelter, werealso discussed. The oceanography of this area was also studied and presented in theproceedings of a workshop on the Kitimat marine enviromnent in 1983 (170). Thecontamination of Kitimat Arm by PAils is well recognized (64-66, 72). Some informationon the concentrations of PAHs in Kitimat Arm sediments is available; however, thedistribution and fate of PAils throughout the Kit.imat Arm is not known. Prior to thisthesis work, no study on the influence of microbial activity on the fate of PAHs in thisparticular environment has been carried out.21Information on the distribution of PAHs in Kitimat water body is very limited (64,72), because of their low concentrations in water. In order to study the distribution ofPAHs in the Kitimat water body, preconcentration treatment is necessary. A recent studyconducted by Environment Canada and Alcan (64) showed that the concentrations of thesixteen PAHs were generally below the detection limit (0.01 nglml), and tetracyclic PAHswere more frequently detected than the low molecular weight PAHs. These resultsreinforce the need for study of microbial transformation of high molecular weight PAHs inthis environment, because these PAHs are potential carcinogens, and living organisms inthe water are in direct contact with these PAHs.1.7. Objectives and overview of the thesisThe overall objective of this thesis is to study the microbial degradation of PAHsby microorganisms obtained from Kitimat Arm. These studies are important not onlybecause some unique PAH-degrading microorganisms may be obtained but also becausethey are useful for addressing important issues such as the fate and persistence of PAHs inthe Kitimat Arm environment, an area known to be highly contaminated by PAHs.Prior to this thesis work, there was no information available about the microbialtransformation of PAHs in the Kitimat Arm environment. Therefore the initial studieswere carried out to obtain PAH-degrading microorganisms from this environment.Phenanthrene was used to enrich the cultures. A series of experiments were conducted tostudy the degradation of phenanthrene by the microorganisms obtained from Kitimat Armsea water and marine sediments. These microorganisms were able to utilize phenanthreneas the sole carbon source. Some important factors affecting the degradation were studied.The isolation and partial characterization of the bacteria were also described (Chapter 2).Pyrene, one of the priority pollutants listed by the US EPA, was determined to bepresent at a high level in sediments from Kitimat Arm and in Alcan lagoons, the principal22point sources of PAHs. Pyrene is often considered recalcitrant to be degraded insediments. Only very limited studies have dealt with microbial degradation of pyrene. Sofar, only few strains capable of utilizing pyrene as sole carbon source was isolated.Therefore, a great deal of effort in this thesis was made to establish cultures and todemonstrate that the microorganisms obtained from Kitimat Arm samples are able toutilize pyrene as sole carbon source. The degradation of pyrene was determined bymonitoring both the loss of pyrene and the production of its metabolites. The identificationof the metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene, in the cultures was studied byusing a variety of analytical techniques. Mineralization of pyrene was also assessed(Chapter 3).Although a few laboratory studies have demonstrated bacterial degradation ofpyrene, little evidence for the microbial degradation of pyrene in the natural environment(in situ degradation) has been reported. In this thesis two approaches were taken in orderto confirm that microbial degradation observed in laboratory studies also occurs in thenatural environment of Kitimat Arm. The 14C labeled pyrene was used to study themineralization of pyrene by indigenous microorganisms. Also, determination andidentification of a specific metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene, from pyrenedegradation were carried out. The presence of this typical metabolite gives strongevidence showing that microbial degradation of pyrene occurs in the natural environmentof Kitimat Arm. A number of factors possibly influencing degradation process were alsostudied and presented in Chapter 4.Kitiniat Arm is an unique environment that is highly contaminated by highmolecular weight PAHs. In order to gain some information about the fate and persistenceof these PAHs, the degradation of benz(a)pyrene, a well known carcinogen, was studied.Preliminary results on the degradation of a mixture of PAHs are also presented (Chapter5).23Nonpolar organic compounds such as PAHs and polychiorinated biphenyls (PCBs)in water are at very low concentrations and difficult to determine quantitatively. A simpleand convenient method for preconcentration is required. Initially disposable cartridgescontaining C18 packing materials were used for the preconcentration of PCBs in watersamples. It was observed that trace amount of organic compounds like PCBs adsorb onthe surface of sample container, particularly polytetrafluoroethylene (PTFE) material.Therefore, a method based on the use of a PTFE tubing for theextraction/preconcentration of PCBs and PAHs from water samples was developed. Itinvolves the passage of the water sample through the PTFE tubing followed by the elutionof the adsorbed compounds by solvent desorption. These results are discussed in Chapter6.24Chapter 2.Degradation of Phenanthrene by MicroorganismsIsolated from Marine Sediment and Sea Water2.1. IntroductionPolycyclic aromatic hydrocarbons (PAHs) are considered to be priority pollutantsby United States Environmental Protection Agency (2) and some are known to bechemical carcinogens and mutagens (2, 171-174). PAHs are ubiquitous in theenvironment (1, 36, 67), and anthropogenic activities have resulted in an increase in theirconcentrations. Incomplete combustion and discharge of industrial wastes often are majorcontributors of PAHs to the environment near urban and industrial sites along with othersources (1, 36, 67, 175-177). Microbial degradation of PAHs has an important influenceon their fate and persistence in a particular environment. This concept has been the basisof a number of studies of the degradation of organic compounds, including PAHs, bynaturally occurring microorganisms (94, 105, 106, 178, 179). A great deal of effort hasbeen directed to the use of indigenous microorganisms to accomplish bioremediation ofsites contaminated by oil spills and other waste (102, 137, 180-182).Kitimat Arm, British Columbia, Canada is an area under environmental study.Over 120 tons of PAHs are produced each year from a nearby aluminum smelter becauseof the use of large amounts of coal tar and pitch in the smelting process. As a result,elevated levels of PAHs are found in the local environment. For example, the PAHconcentration in near-shore sediment samples in Kitimat harbor is up to 528 p.g g1 (64-66). The objective of this present study is to establish if PAH-degrading microorganismsare present in this particular environment. This forms the first part of a series of studiesinvolving an assessment of the environmental fate of PAHs in Kitimat Arm.PAH-contaminated sea water and marine sediment samples obtained fromKitimat Arm were used as initial inoculants in order to enrich specific microorganisms25for use in degradation studies. Phenanthrene and pyrene, among the 16 priority PAHpollutants named by the U.S. EPA were chosen as examples for the study because theyhave intermediate solubility and volatility. Furthermore they are among the mostabundant PAHs found in the study area (64, 65).2.2. Experimental2.2.1. Instrument and AnalysisA high performance liquid chromatography (HPLC) system, consisting of aWaters 600E multisolvent delivery unit, a system controller, and a U6K sample injector,equipped with a UV/visible spectrophotometer (Waters Lambda-Max Model 481 LCSpectrophotometer) and a scanning fluorescence detector (Waters Model 470) was usedfor the determination of phenanthrene. A reversed phase C18 column (Supelcosiltm’LCPAH, 25 cm x 4.6 mm i.d., 5 tm particle size, from Supelco, Inc.) with a 3-cm guardcolumn of the same packing material was used. Isocratic HPLC runs were performed byusing acetonitrile/water (70/30) as eluent and the flow rate was kept at 1.0 mi/mm. Priorto the HPLC analysis, all the culture samples were centrifuged on a Dynac centrifuge(Becton Dickinson Co.) at its full speed for 25 mm. A 20 tl aliquot of a sample was usedfor the HPLC analysis. The retention times for phenanthrene and pyrene were 8.4 mmand 12.6 mm, respectively.The wavelength of the UV/visible spectrophotometer was set at 254 nm. Theoptimum excitation and emission wavelengths for the determination of phenanthrene byusing fluorescence have been reported (158, 183) to be 252 and 370 nm, respectively.These wavelengths were used in the present study. An excitation wavelength of 333 nmand emission wavelength of 373 nm (183) were used for the determination of pyrene.During some initial experiments, a Shimadzu UV-2100 UV/visiblespectrophotometer with wavelength scanning capability was also used to monitor anychange in the composition of culture medium.262.2.2. MaterialsPhenanthrene were obtained from Aldrich (Miowaukee, WI). HPLC gradeacetonitrile and acetone were obtained from Fisher Scientific.Mineral salt (M.S.) medium: The mineral salt medium was the same as thatreported by Mueller et al. (184). It consisted of (mgfL) (NH42S0 (1000), K2HPO4(800), KH2PO4 (200), MgSO47H2O(200), CaC12HO(100), FeC136H2O(5), and(NH46Mo7O24 (1). It was sterilized (Amsco, Eagle Series-3041 gravityautoclave) before the addition of PAH for degradation studies. The pH of the mediumwas 7.2.M.S. medium containing phenanthrene-acetone (medium A): A known amount ofphenanthrene was added as an aliquot (0.1 ml of acetone solution) to the sterile M.S.medium. The final concentration. of phenanthrene was 5-10 igIm1. The phenanthreneacetone solution was found to be sterile: no growth of bacteria was observed over twomonths when a phenanthrene-acetone solution was plated on a half strength marine brothagar (Difco 2116) plate.M.S. medium containing phenanthrene and Tween-80 (medium B): excess solidphenanthrene was added to M.S. medium containing 200 mgIL of Tween-80. Themixture was stirred overnight to dissolve as much solid phenanthrene as possible,autoclaved, and then filtered through sterile glass wool. The filtrate was used as theculture medium, and the final concentration of phenanthrene in this medium wasdetermined by using HPLC/UV absorption spectrophotometry. The addition of Tween80, a nonionic surfactant, served to solubilize phenanthrene and to stabilize the medium.Phenanthrene was not degraded in uninoculated controls containing Tween-80 [27].M.S. medium with solid phenanthrene (medium C): Solid phenanthrene wasadded at a dose of 5 mg per 50 ml of M.S. medium. The mixture was autoclaved beforeuse.272.2.3. CulturesAll the sea water and marine sediment samples were obtained, near shore, asshown in Figure l.5b. Sea water samples were collected by dipping sterile evergreentubes (60 ml) into the water, and about 30 ml of each sample was directly inoculated intoa sterilized dilution bottle (160 ml) containing 50 ml of M.S. medium and 5 mg of solidphenanthrene (medium C). Surface sediments at the top 10 cm were obtained by grabsampling. The original sediment cultures were initiated by inoculating approximately 2 gsediment into 50 ml of medium C. In the same way, replicate cultures were obtained byinoculating sea water or sediment in M.S. medium without the addition of PAH. Thelatter cultures were used to study whether or not PAH-degrading microorganisms areavailable without further PAH induction in the enrichment process. These cultures werebrought back from the field to the laboratory within 24 hr. The cultures were kept on ashaker (136 RPM) at 26 °C. Two weeks later, 5 ml aliquots of each culture were dilutedwith 50 ml of medium C to obtain subcultures.After each sea water culture was incubated for one week, 5 ml aliquots wereagain transferred to 50 ml of medium C. After two more transfers, three types of culturesI, II, and III were obtained by separately inoculating the subculture into media A; B; and,C, respectively. These three types of cultures were used for the degradation studies. Thecultures were maintained by transferring them (1:10) with the corresponding mediumevery two weeks.Sediment cultures were enriched by transferring 5 ml aliquots of the originalcultures into 50 ml of fresh medium C. When phenanthrene particles disappeared fromthe solution approximately 1.5 months after the transfer, a further similar transfer wasconducted. After a total of three such transfers and enrichment processes (total incubationtime of six months), the enriched cultures (i.e. the third subcultures) were used fordegradation studies.28Isolation and purification of strains were carried out on half strength marinebroth (Difco 2116) agar plates. Four strains were obtained from a sea water cultureenriched in medium A (10 pgIml), and they were allowed to grow on half strengthmarine broth slants for one week before being transferred separately into fresh mediumA. The resulting four pure cultures were then used for further degradation studies.2.2.4. Degradation ExperimentErlenmeyer flasks, pipettes, and culture media were all sterilized. An aliquot of10 ml of M.S. medium was pipetted into each sterile 50 ml Erlenmeyer flask, to which0.1 ml of phenanthrene (1000 or 500 pgIml) in acetone was added. The desired amountof culture (0.1-1 ml) was added to each flask. The flasks were kept on a shaker at 26 °C.All these procedures were done under sterile conditions in a laminar flow hood. Parallelsterile controls were prepared similarly except that no culture was added. Killed cellcontrols were obtained by autoclaving the flask containing the added culture.Triplicate cultures and controls were amended with 10 ml of acetonitrile (HPLCgrade, Fisher Scientific) at several intervals of incubation time to stop growth and tosolubilize the PAH. The flasks were placed back on the shaker and shaken overnight. Thesamples were centrifuged and supernatants were analyzed by using HPLC with UV andfluorescence detection. Recoveries of phenanthrene in the controls were 90-100%.In another set of experiments, phenanthrene (10 .tg in 0.1 ml of acetone) wasadded into each sterile 50-ml Erlenmeyer flask. The acetone was allowed to evaporatecompletely by leaving the flask in a fumehood overnight. M.S. medium (10 ml) and anappropriate aliquot (0.1-1 ml) of culture was then added to the flask for the degradationstudy. Phenanthrene was added as the sole carbon source in this experiment.For each experiment, the same volume of a particular culture was added to eachflask. The initial amounts of cells introduced to each flask were assumed to be the same,although no biomass measurement was carried out.292.3. Results and Discussion2.3.1. Degradation of phenanthrene by microorganisms enriched from sea waterInitial studies on the degradation of phenanthrene were carried out by inoculatingan enriched sea water culture into medium A (phenanthrene in acetone). The absorptionspectrum of the supernatant of the culture was monitored as a function of incubation timeafter acetonitrile was added to the culture to stop growth. Figure 2.1 shows the UVspectra of the supernatants of the culture at incubation time 0, 10, and 20 hr after 10 mlof medium A (containing 10 ig ml1 of phenanthrene) was inoculated with 1 ml ofculture I. The concentration of phenanthrene in the culture was reduced to half within 10hr of incubation. Very little phenanthrene remained after 20 hr of incubation. Theconcentration of phenanthrene in sterile controls remained unchanged throughout thisexperiment, indicating that the disappearance of phenanthrene did not occur in theabsence of microbial activity. These results clearly show that phenanthrene is rapidlydegraded by the microorganisms. The appearance of a wide shoulder between 260 and300 nm in spectrum (c) of Figure 2.1 is possibly associated with metabolites ofphenanthrene degradation.In the light of these results, further studies on the microbial degradation ofphenanthrene were carried out making use of HPLC with UV absorption andfluorescence detection to quantitatively determine the concentration of phenanthrene inthe culture. Three types of cultures, I, II, and III were initiated from sea water samples byenriching microorganisms in media A (phenanthrene-acetone), B (phenanthrene-Tween80), and C (phenanthrene alone), respectively. These enriched cultures were used asinoculants to study the degradation of phenanthrene in medium A. The results aresummarized in Table 2.1.30F)4-00000I-.0001)4-00F’)00000000500 o0I000 2000240 300 360 420Wavelength, nmFigure 2.1. Absorption spectra of the seawater culture: 10 ml of medium A inoculated with1 ml of culture I at incubation time (a) 0, (b) 10 hr, and (c) 20 hr.31Table 2.1. Degradation of phenanthrene by three cultures I, II, and IIICulture* Phenanthrene Medium Culture Completein Culture (ml) Inoculated (ml) Degradation TimeCulture I Medium A (phenanthrene-acetone)10 pg/mi 10 0.1 23 hr6pg/ml 10 0.1 20hr8pg/ml 10 1 lOhr0.2 pg/mi 10 0.1 6 hr0.2 pg/mi 10 1 1 hrCulture II Medium B (phenanthrene-Tween 80)10 pg/mi 10 2 2OhrCulture III Medium C (phenanthrene alone-mostly undissolved)10mg 100 6 50 days1mg 100 6 8days* Described in the experimental section.When 10 ml of medium A (containing 10 pg/mi of phenanthrene) is inoculatedwith 0.1 ml of culture I, the time required for complete degradation (below the detectionlimit of 1 ng/ml) was 23 hr. By increasing the amount of culture I from 0.1 ml to 1 ml,this time was reduced to 10 hr. Complete degradation of phenanthrene at a reducedconcentration (0.2 pg/mi) in medium A took only 6 hr with 0.1 ml of the culture and 1 hrwith 1 ml of the culture I. When 10 ml of the medium B was inoculated with 2 ml ofculture II, over 96% of the phenanthrene was degraded after 20 hr of incubation. Theseresults demonstrate that microorganisms originating from sea water samples from32Kitimat Arm are able to degrade phenanthrene rapidly. Table 2.1 also shows thatphenanthrene introduced as solid can also be degraded. When 6 ml aliquots of culture IIIwere added to 100 ml of medium C, the phenanthrene particles (10 mg) disappeared after50 days of incubation. The disappearance of 1 mg of phenanthrene particles wasobserved after 8 days of incubation. The fact that phenanthrene is added as solid withoutany accompanying organic solvent indicates that this mixed culture may be capable ofusing phenanthrene as the sole carbon source. Microbial isolates capable of utilizingphenanthrene as sole carbon sources have been reported by others (94).2.3.2. Degradation of phenanthrene by microorganisms enriched from sedimentMicroorganisms were enriched from two sediment samples by using medium C,phenanthrene being the sole carbon source. After three subsequent transfers and a total ofsix months of incubation at 26 °C, both enriched sediment cultures were able to degradephenanthrene. Phenanthrene concentration in the culture was quantified by usingHPLC/fluorescence. Phenanthrene was completely degraded within two days after 1jig/ml of phenanthrene in 10 ml M.S. medium was inoculated with 0.5 ml of the enrichedculture.Two cultures from sediment samples that had been enriched in mineral saltmedium only, i.e. no phenanthrene was added in the enrichment process, were also usedto study phenanthrene degradation. After four days of incubation, one culture showedphenanthrene degradation and the other did not. When these two sediment cultures wereinoculated on half strength marine broth agar plates, the culture which degradedphenanthrene showed growth within two days while the other culture only showedgrowth after nine days of incubation. This suggests that more viable organisms werepresent in the phenanthren-degrading cultures. The sediment cultures enriched withphenanthrene as discussed earlier also showed growth of microorganisms on the agarplate within two days of incubation.33These results indicate that phenanthrene-degrading microorganisms are present inthe Kitimat Arm sediment samples. The ability of these organisms to degradephenanthrene may result from their previous exposure to this PAH in the sediment. Theconcentration of phenanthrene in the sediment sample was found to be approximately 13igIg (dry weight) (65).2.3.3. Isolation and biolog tests of strainsThe degradation of phenanthrene described above was carried out by using amixed community of microorganisms enriched from sea water and sediment. Furtherstudies on the degradation of phenanthrene by pure cultures were also carried out. Forthis purpose, a sea water culture enriched in medium A, was isolated on half strengthmarine broth agar plates. Four strains were obtained and purified. These strains werepartially characterized by using biolog comparative metabolite analysis tests (185, 186)in Department of Microbiology at the University of British Columbia, and the results aresummarized in Table 2.2. The microorganisms are all Gram stain test negative and rodshaped. Comparison of the biolog number with those in the bio-data bank reveals thattwo of the microorganisms were tentatively identified as Moraxella atlantae andAlteromonas haloplanktis, respectively. The other two are not recorded in the data bankbut they appear to have a similarity to the Comamonas genus for strain Y, and theEnterobacter genus for strain LY.Each of the four isolated strains was incubated separately in the medium A, andthe ability of these pure cultures to degrade phenanthrene was then studied. It was foundthat one of the unidentified strains (LY) degraded phenanthrene the fastest. Phenanthrene(10 igIml) in 10 ml of medium A was degraded to an undetectable level within 22 hrwhen 0.2 ml of the LY pure culture was inoculated into the medium. The otherunidentified strain (Y), with yellow color, was also able to utilize phenanthrene but at amuch slower rate. The remaining two identified strains did not degrade phenanthrene34after 11 days of incubation, however, it is not known if these strains have an impact onthe degradation rate when they co-occur with other microorganisms. The HPLCchromatogram with fluorescence detection obtained from the single strain culture (LY) issimilar to that from the mixed culture. Thus, it is possible that strain LY is the mainmicroorganism degrading phenanthrene.Table 2.2. Biolog tests of four stains isolated from a sea water cultureStrain Shape Gram Stain Biolog Number Match& Size Test* DataBankW rod - 0004-0000- 1000-0450- Moraxellal.0x2.0 pm 0104-0000-0040-0000 atlantaeP rod - 3400-1304-3555-6270- Alteromonas0.5x1.0 lIm 0570-0177-1114-0000 haloplanktisY rod - 0300-0000-1002-4542- no match, similar tol.0x2.0 im 0175-6434-5144-6000 Comamonas genusLY rod- 3660-7775-7776-7777- no match, similar to0.4x 1.0 im 2575-7777-7776-3436 Enterobacter genus* indicates Gram stain test negative.2.3.4. Factors influencing phenanthrene degradationSome of the factors that may affect the degradation rate of phenanthrene werefurther studied by using a sea water culture (culture I) as the inoculant. It was found thatthe rate of phenanthrene degradation was dependent on the volume of the inoculant. Thedegradation occurs in two phases as seen in Figure 2.2. In the initial phase, phenanthrenewas degraded at a relatively slow rate; whereas in the second phase, the degradation wasrapid. When 0.1 ml of the mixed culture was inoculated into the medium containing35either 10 .tg/mi or 6 IgIml of phenanthrene, the period of time of the initial phase wasapproximately 10 hr. This initial period was reduced to 5 hr when 1 ml of the mixedculture was used. The time required for the complete degradation of phenanthrene wasalso reduced to half when the amount of the culture is increased from 0.1 ml to 1 ml. Theresult that the inoculant size affect the rate of phenanthrene degradation is in goodagreement with those reported by others (105, 127).IFigure 2.2. Effect of amount of bacteria on the degradation of phenanthrene by sea water culture I(•) 1.0 ml, and (A, R)0.1 ml of culturel(0) 0.1 ml of autoclaved culture I(Q) sterile control without culture4200 10 20 30 40Incubation Time, hr5036The effect of phenanthrene concentration on the degradation rate was alsostudied. When 0.1 ml of culture I was separately inoculated into the medium containing10, 6, and 0.2 tg/ml of phenanthrene , the degradation rates during the initial phase, interms of loss of phenanthrene per hour, are 0.22, 0.12, and 0.04 pg ml per hour,respectively. Thus the higher the concentration of phenanthrene present in the medium,the faster the initial degradation rate was. It has been reported that the rates of PAHsmineralized to CO2 and H20 by microorganisms are directly related to PAHconcentrations (127, 184, 187-189) and are controlled by the extent of the pollutantloading and/or pre-exposure (187, 189-191). The present results are in good agreementwith these findings. In addition, these results show that the time required for thecomplete degradation of phenanthrene is dependent both on the population of themicroorganisms and on the concentration of phenanthrene.Nutrients can have considerable effect on the growth of microorganisms (192);therefore, the effect of the culture medium composition on the degradation ofphenanthrene by the mixed microorganisms was studied by eliminating one component(Fe3+, Mo(VI), phosphate, NH4+, or SO42-) at a time from the mineral salt medium. Itwas found that the elimination of Fe3+ (5 jig/ml) from the medium results in a decreasein degradation rate. As shown in Table 2.3, after 29 hr of incubation over 99% of theinitial phenanthrene was degraded in the medium containing all the ingredients; whereasonly 30% of the phenanthrene was degraded in the same time when Fe3+ was absentfrom the medium. The elimination of other components had little effect. When sterilenatural sea water was used as medium instead of mineral salt, degradation was notobserved after 40 hr of incubation, and the addition of Fe3+ into the sea water did notenhance the rate. The inhibition of the degradation in sea water was probably due to thepresence of high concentration (3%) of NaC1, suggesting that this culture would bedifficult to grow in the natural marine environmet. This contention is supported by thefinding (Table 2.3) that no degradation is detected when 3% NaC1 is present in the37mineral salt medium. The elevated NaC1 concentration probably causes an increase inosmotic pressure, resulting in the inhibition of growth of the microorganisms (192).Table 2.3. Effect of Fe3+and NaC1 on degradation of phenanthrene by sea water cultureIMedium Percent of Initial Phenanthrene (%) at Incubation Time2Ohr 29hr 4OhrControl 101± 4 x 92±3M.S. medium 67 ± 11 0.6±0.1 ndFe3 absent 80±7 72±6 9±2Seawater x x 94±3Sea water +FeC13(5mgL1) x x 101±4M.S. medium +3%NaCl x x 99±4x -- not evaluated.nd -- no phenanthrene detected.38Some common carbon sources such as glucose, peptone and yeast extract wereadded to the medium to examine if they increase the ability of the microorganisms totolerate saline condition. The results of the degradation of phenanthrene obtained fromthe cultures in mineral salt medium, and in mineral salt medium with the addition ofNaCl, glucose, peptone and yeast extract are summarized in Figure 2.3. The relativeconcentration of phenanthrene was obtained by comparing the concentration ofphenanthrene in the cultures with that in sterile controls after the same incubation time.The results show that phenanthrene was completely degraded after 23 hr in M.S. mediumand after 30 hr in M.S. medium containing additional 1% NaC1. When 3% NaCl waspresent in the M.S. medium, phenanthrene was not degraded within 170 hr. The additionof 250 ig m11- glucose to the medium did not increase the extent of degradation in thesame incubation period. However, when peptone and yeast extract were added to theM.S. medium containing 3% NaC1, the rate of degradation was increased significantly,and phenanthrene was completely degraded after 170 hr. The addition of yeast extract (avitamin source) and peptone (a nitroge source) probably benefits some nutritionallystressed microorganisms, resulting in a higher population of microorganisms in theculture. As a consequence of the increased microorganism population, the degradation isalso enhanced.3960I_______ ________•0 50 100 150 200Incubation Time, hrFigure 2.3. Effect of culture medium on the degradation of phenanthrene by sea water culture I(•) M.S. medium(0) M.S. medium + 1% NaCI(D) M.S. medium +3% NaCI(1) M.S. medium + 3% NaCJ + 250 .1.g m11 of glucose(A) M.S. medium +3% NaC1 + 250 p.g m11 of peptone and yeast extract40Chapter 3.Degradation of Pyrene and Characterization of Metabolites3.1. IntroductionIt is demonstrated in Chapter 2 that microorganisms, which can degradephenanthrene, are present in sea water and sediment samples obtained from Kitimat Ann(193). The fast degradation of phenanthrene by these indigenous microorganismsprompted further studies on the microbial degradation of other PAHs.Microbial degradation of low molecular weight PAHs, such as naphthalene (194-198), phenanthrene (119, 198-202), and anthracene (119, 202) has been proven to berelatively facile. These PAHs can be metabolized or co-metabolized with otherhydrocarbons by using either mixed microbes or pure microorganisms (94, 105, 203,190). However, PAHs with four or more fused aromatic rings are more recalcitrant tomicrobial attack, and axe degraded only with difficulty (102, 106, 182). The persistenceof PAHs in the environment increases with the number of aromatic rings in the PAR Asa consequence, while degradation of low molecular weight PAHs has been studiedextensively (94, 102, 105, 106, 182), only few examples have been reported of the cometabolism (101, 204-206) and metabolism (100, 207, 208) of PAHs having four ormore fused aromatic rings. The microbial degradation of these higher molecular weightPAHs has attracted much attention recently (102, 106).Pyrene, a PAH composed of four fused benzene rings, which is widespread in theenvironment of Kitimat Arm at considerable concentrations, up to 40 tg/g in sedimentsamples (66), merits a detailed microbial degradation study. This environment shouldprovide possible sources of microorganisms which have been exposed to a highconcentration of pyrene, and the pre-exposure to pyrene should contribute to their abilityto degrade this PAH, because it is generally believed (123, 127, 129, 191, 209, 210) that41the enzymatic capability of microorganisms to degrade certain pollutants may depend onexposure to levels of the pollutant sufficient to induce the necessary degradative enzymein the microorganisms, and the population of PAH-degrading microorganisms may beenriched. Therefore, it was decided to study the degradation of pyrene by using microbesobtained from water and sediment samples of the Kitimat Arm environment.In order to understand the degradation process, it was also necessary to determinethe metabolic product of the degradation.3.2. Experimental3.2.1. InstrumentationThe instruments used in these studies included a scanning UV/visiblespectrophotometer and a HPLC coupled to UV and fluorescence detection, as describedin Section 2.2.1. Two analytical reversed phase HPLC columns (Supelcosil LC-PAH,250 mm x 4.6 mm,; and GL Scientific ODS-2, 250 mm x 4.6 mm, 5im) and apreparative HPLC column (Waters RCM 100 mm x 25 mm) were used for the separationof pyrene and metabolites. A Packard 1900 TR Liquid Scintillation Analyzer was used tomeasure the radioactivity of 14C in labeled pyrene metabolites and the 14C02 evolvedfrom the mineralization of pyrene.A GC/FID system (HP 5890) and a GCIMS system (VG 7070E) was used toanalyze the acetylation products of the standard, cis-4,5-dihydroxy-4,5-dihydropyrene,and the metabolites from pyrene degradation. A capillary column, DB-5 (30 m x 0.25mm i.d., 0.25pm) was used in both systems to separate the components.3.2.2. Chemicals and culture mediaBoth unlabeled and the 14C labeled pyrene standards were obtained from Sigma(St. Louis, MO). Cis-4,5-dihydroxy-4,5-dihydropyrene was a gift from Dr. C. E.42Cerniglia of the National Center for Toxicological Research, Jefferson, Arkansas. [-4C-4,5,9,10]-pyrene was supplied as a toluene solution (52 p1) with a specific radioactivityof 40-60 mCilmmole (1 pCiJp.l). The ‘4C radioisotope produces 13 radiation (0.156Mev) and has a half-life (t112) of 5730 years. Appropriate care was taken to meetradioactive safety requirements in handling this material. Phenanthrene,benzoanthracene, ethanol amine, and ethylene glycol monomethyl ether were obtainedfrom Aldrich. The liquid scintillation cocktail (LSC) containing a mixture of solvent,primary fluor, and secondary fluor, was obtained from Fisher Scientific, and was used forthe liquid scintillation counting experiment. Marine broth and casitone were obtainedfrom Bacto. The CGY medium (pH 7) (also from Bacto) consisted of casitone (5 gIL),glycerol (10 milL), and yeast extract (lg/L).The same mineral salt (M.S.) medium as described in Section 2.2.2 was usedthroughout this study. The culture enrichment medium was prepared as follows. Thesolid PAH (5 mg of phenanthrene, pyrene, or benzoanthracene), was added to 50 ml ofM.S. medium and autoclaved prior to use. The media so prepared were used for theenrichment of sea water and sediment cultures. No other source of carbon or energy wasadded to these media.3.2.3. CulturesOriginal cultures were prepared by inoculating approximately 2 g of sediment or30 ml of sea water obtained from near shore of the Yacht Club in Kitimat Arm (Figurel.5b) into 50 ml of one of the three enrichment media in a 160-ml dilution bottle.Duplicate cultures were prepared by using the same enrichment medium and a sampleobtained at the same location. Thus six sea water cultures and six sediment cultures wereobtained, containing added phenanthrene, pyrene, or benzoanthracene. Duplicate seawater and sediment “reference” cultures were prepared similarly by inoculating sea wateror sediment (the same amount as above) into 50 ml of mineral salt medium but without43adding any PAH. These four “reference” cultures were used to investigate if additionalPAH induction is necessary in order to degrade pyrene. The compositions of the originalcultures is summarized in Table 3.1. These sixteen cultures were kept in the dilutionbottles and were used as inoculants for further enrichment.NameTable 3.1. Composition of original cultures*PAH added Sample addedReference sediment-iReference sediment-2Reference water-iReference water-2Phenanthrene sediment-iPhenanthrene sediment-2Phenanthrene water-iPhenanthrene water-2Pyrene sediment-iPyrene sediment-2Pyrene water-iPyrene water-2Benzoanthracene sediment-iBenzoanthracene Sediment-2Benzoanthracene water-iBenzoanthracene water-2nononono5 mg phenanthrene5 mg phenanthrene5 mg phenanthrene5 mg phenanthrene5 mg pyrene5 mg pyrene5 mg pyrene5 mg pyrene5 mg benzoanthracene5 mg benzoanthracene5 mg benzoanthracene5 mg benzoanthracene2 g sediment2 g sediment30 ml sea water30 ml sea water2 g sediment2 g sediment30 ml sea water30 ml sea water2 g sediment2 g sediment30 ml sea water30 ml sea water2 g sediment2 g sediment30 ml sea water30 ml sea water* All cultures contain 50 ml of mineral salt medium.44Enrichment cultures were obtained by transferring 5 ml of the original cultureinto 50 ml of fresh enrichment medium containing mineral salts and the appropriatePAll. After approximately six weeks, the cultures were further transferred into the freshmedium to selectively enrich the PAll-degrading microorganisms. Following threefurther subsequent transfers, the enriched cultures so obtained were used for degradationstudies. These enriched cultures were maintained by transferring the cultures into freshenrichment medium every six weeks at a dilution factor of 1:10. The reference symbolsattached to the enriched cultures are derived from the original cultures. For example,Pysed-1 originated from the Pyrene sediment-i culture.3.2.4. Preservation of culturesPreservation of some pyrene-degrading microorganisms was carried out in orderto preserve the microorganisms for longer term and for later access. One ioop of eachculture (approximately 0.1 ml) was streaked on a slant containing half-strength marinebroth and agar. The slant was kept at 26 °C for a week in order to allow themicroorganisms to grow on it. The sterile enrichment medium (1.0 ml) with appropriatePAH (0.2 mg/ml) and glycerol (20%, 1.0 ml) was then added to each slant. Themicroorganisms grown on the slant were scratched off and mixed with the medium byusing a sterile ioop. The mixture was then transferred into two sterile cryogenic vials.These vials were first put in dry ice for 45 mm and then placed in liquid nitrogen for longterm storage. A few of these preserved cultures were tested for their viability and theirability to degrade pyrene after one and seven months of storage, respectively. Similarresults of pyrene degradation were obtained by using these preserved cultures ascompared to the fresh cultures, indicating that the preservation of the cultures wassuccessful.453.2.5. Isolation an,d biolog test of strainsThe isolation of cultures was carried out in order to obtain and characterize purestrains that were capable of degrading pyrene. Three enriched cultures including Pysed-1,Pysed-2, and Pyw-1 were each diluted by dilution factors of 100, 1000, and 10,000 usingsterile mineral salt medium containing pyrene (0.1 mg/mi). Another enriched culture,Phensed-2, was diluted similarly by using mineral salt medium containing phenanthrene.An aliquot (0.1 ml) of each diluted culture was plated on a half strength marine broth agarplate. After 2 days of incubation under the temperature 26 °C, the growth of colonies wasobserved from all of the plates except from Pyw-1 at dilution factor of 10,000. No colonywas observed on sterile control plates. When the incubation time was extended to 7 days,colonies grown from Pyw- 1 were also observed. Two distinct colonies with light yellowand yellow color were observed from each agar plate from the 10,000 times dilutedcultures. Both colonies were spherical in shape and the light yellow (ly) colonies wereslightly bigger than the yellow (y) colonies. These two types of colonies were furtherpurified by transferring each separately to freshly prepared agar plates. After threesubsequent such transfers, the colonies were grown on half strength marine broth slantsfor a week, and then transferred to liquid enrichment medium. Thus eight cultures,namely Pysed-1-y, Pysed-1-ly, Pysed-2-y, Pysed-2-ly, Pyw-1-y, Pyw-1-ly Phensed-2-y,and Phensed-2-ly were obtained from the above four enriched cultures; each enrichedculture resulted in two cultures.The eight cultures were sent to the Microbiology Department at UBC for biologtests. Each of the strains was re-streaked onto CGY medium for further purification.While most were found to contain uniform colonies, each of the strain Pysed-l-y, Pyw-1-y, and Phensed-2-Iy resolved into two kinds of colonies. Therefore, the latter three strainswere further purified on the CGY media to produce six isolates. Thus a total of 11isolates were obtained from the four enriched cultures. These 11 isolates were partially46characterized by using the Gram stain test and the Biolog comparative metabolicanalysis. The results of these tests are summarized in Table 3.2.Table 3.2. A summary of partial characterization of 11 isolatesStrain Shape Gram test Biolog # Match data bank Correlation& Size(pm)Pysed-1-ly Curved rod - 0000-0000-0003-6406 Bordetella good0.8x5.0- 1501-6577-0100-0000 parapertusisPysed-1-y Swollen rod - 1701-0725-2202-0042 Pseudomonas poor(small) 0. 8x 1.0-2.0 - 1304-4037-3104-0000 paucimobilisPysed-1-y Swollen rod + nd nd nd(big) 2.0x2.5-6.0Pysed-2-ly Rod - 0304-0000-0003-7406 Alcaligenes good1 .0x2.0-5.0 -7527-6774-5754-4100 faecalisPysed-2-y Encapsulated - 0300-0000-0023-0002 Moraxella goodrod 1 .0x2.0-3.0 -0002-0004-0000-0000 PhenylpyruvicaPyw-1-ly Encapsulated 3504-3305-1002-0002 Pseudomonas poorrod 0.7x3.5 0001-0475-0100-0002 vesicularisPyw-1-y Rod - 0300-0000-0023-4002 Moraxella poor(small) 0.5x3 .0-7.0 -1023-4017-0100-4000 bovisPyw-1-y Rod- 0301-0000-0023-4002 Moraxella poor(big) 0.4x 10-30 0003-4417-0100-4000 bovisPhensed-2-ly Rod - 0026-3503-1302-4402 Pseudomonas poor(white) 0. 8x2.0-4.0 0120-0124-7516-2000 genusPhensed-2-ly Rod - nd nd nd(ghost) 1x2.0-6.0Phensed-2-y Encapsulated 17 12-2304-2072-0402 Pseudomonas poorrod 0.7x2.0 1000-0015-1104-1003 vesicularisnd- not determined47The light yellow strain (Pysed-1-y, big) that was resolved from Pysed-1-y onCOY media after isolation had poor growth, and the Biolog test was not able to becompleted. Similarly, isolate Phensed-2-y (ghost) grew poorly after the purification.3.2.6. Measurement of culture growthGrowth of the selected cultures including mixed culture Pysed-l, Pysed-1-y, andPyw-1-y, as examples, was monitored by determining optical density of the cultures at500 nm wavelength. Growth curves were obtained by plotting the optical density versusthe incubation time, as shown in Figure 3.1. No growth was observed in the control.Little growth was determined from the measurement of optical density when 20 ig ofpyrene, 20 ml of mineral salt medium, and 0.5 ml of Pysed-1 were incubated for 180 hr.This result reflects the low sensitivity of the optical density measurement, becausepyrene-degrading microbes did grow during this period of time as judged from pyrenedegradation studies that will be described below. However, the addition of yeast extractsignificantly enhanced the growth of microorganisms in the culture. Thus a rapidincrease in the optical density of the culture was observed after 28 hr of incubation whenyeast extract (250 tgIml) was present in the culture. The maximum growth was reachedat 40 hr of incubation. Similar results were obtained when the growth of purified culturePyw-1-y was monitored. The addition of yeast extract also resulted in a faster growth ofmicroorganisms in these cultures (Figure 3.1).480.180.)0-0.02200Incubation Time, hrFigure 3.1. Growth curves of cultures as measured by the optical density of the culture.(0 ) control, mineral salt medium(• ) culture Pysed-1, mineral salt medium(R ) culture Pyw-l-y, mineral salt medium and yeast extract(A ) culture Pysed-1-y, mineral salt medium and yeast extract(D ) culture Pysed- 1, mineral salt medium and yeast extract0 50 100 150493.2.7. Degradation of pyreneAn 100 jtl aliquot of a 100 jig/mi solution of pyrene in acetone was added to thebottom of a sterilized Erlenmeyer flask (50 ml). Presumably, acetone evaporatescompletely after leaving Erlenmeyer flasks with cotton plugs in a sterile Laminar flowhood overnight. In this way, the desired amount of pyrene (10 jig) was quantitativelyintroduced without adding solvent which can be an alternative carbon source formicroorganisms. Sterile mineral salt medium (10 ml) was then quantitatively pipettedinto the flask followed by 0.1 ml of one of the cultures. Sterile controls were preparedsimilarly without adding the culture. These cultures were placed on a rotary shaker (136rpm) for incubation at 26 °C. After a period of incubation time, acetonitrile (10 ml) wasadded to the flask to stop growth. The flasks were placed on the shaker and shakenovernight. The mixture in the flask was centrifuged to remove any suspended particles,and the supernatant was subjected to HPLC analysis for pyrene and metabolites. Sterilecontrols were analyzed similarly to ensure no abiotic loss of pyrene and no production ofmetabolites.When the preserved cultures were used in the degradation studies, the frozenculture (in liquid nitrogen) was allowed to defrost in ambient temperature and was grownin the fresh enrichment medium for two weeks before being used as the inoculant.3.2.8. Mineralization of pyreneThe mineralization of pyrene was studied by monitoring the 14C02evolved fromthe cultures as a result of the mineralization of‘4C-4,5,9,10-labeled pyrene (chemicalpurity 99% and radiochemical purity >95%). One set of experiments was designed todetermine the amount of pyrene mineralized, and the metabolites present, after a suitableincubation time. Another set of experiments was designed to obtain mineralization aftervarious incubation times. For the first set of experiments, 0.1 ml aliquots of pyrenesolution (600 jig/mi) in acetone and 20 jil of 14C labeled pyrene (10 nCi/jil) in50acetone/toluene were added to a sterile Erlenmeyer flask (125 ml). The solvent wasallowed to evaporate by leaving the flasks (fitted with cotton plugs) in a fumehoodovernight. Mineral salt medium (20 ml) was aseptically added to each flask followed by aninoculation of 1 ml of the enriched cultures Pysed-1 or Pysed-1-y. The flask wasimmediately sealed with a sterile serum rubber stopper and covered with aluminum foil.Three flasks containing each enriched cultures and duplicate controls without cultureinoculants were prepared similarly. These cultures were placed on a rotary shaker (136rpm) for incubation. The total amount of 14C02 evolved was determined after 12 days ofincubation.For the measurement of 14C02,an apparatus similar to that described by Fedoraket at. (211) was used for the trapping of CO2. It consisted of a rotometer to control theflow of flushing nitrogen gas, a 25-mi glass vial containing 4 ml of 8 M sulfuric acid andthe injected culture sample consisting of 5 ml liquid and 10 ml of head space gas, and three7-mi liquid scintillation counting vials. Each vial contained a mixture of 0.67 ml ethanolamine, 0.33 ml ethylene giycol monomethyi ether, and 4 ml of liquid scintillation cocktail,and was sealed with a serum rubber stopper. The vials were inter-connected by usingpolyethylene tubing and needles inserted through the rubber stoppers as shown in Figure3.2. A flow of nitrogen (approximately 50 mi/min) was used to purge the 14C02from theacidified sample solution for 10 min, and the displaced gases were bubbled through thetrapping solution. The 14C02was trapped in the vials which were subsequently placed ina liquid scintillation counter for the measurement of radioactivity. It was found thatessentially all the 14CO2 was trapped in the first vial, in agreement with Fedorak et at.(211), who used a commercial CO2 absorbent as trap. The method was used to measurethe 14CO2produced in two flasks from each of the two cultures.51N2To ventFigure 3.2. Apparatus for trapping‘4C02A - flow meter, B - 22 gauge x 7 cm needles, C - 26 gauge x 4 cm needles,D - glass vial (25 ml), E - polyethylene tubing, F - rubber stoppers,1, 2, and 3 - liquid scientillation vials (7 ml).A D 1 2 352The third flask of each batch was measured for 14C-labeled pyrene and ‘4C-metabolites as follows. The whole culture was acidified with 10 ml of 8 M H2S04through the serum stopper and mixed well to release ‘4C02from the culture to the headspace. The flask was purged by introducing a stream of N2 (about 100 mI/mm) for abouttwo hours. The 14C02 in the flask was flushed out and trapped in two vials containing 4ml of a mixture of ethanol amine and ethylene glycol monomethyl ether (2:1) and disposedof properly. After the 1-4C02 was completely flushed out, the culture residue wasextracted with CH21 (10 ml) and the organic layer was separated from the aqueouslayer. One ml of each layer was added to a liquid scintillation sample vial containing 3 mlof the liquid scintillation cocktail. The 14C activity in both organic and aqueous layerswas determined.In addition to the measurement of 14C activity, the pyrene content in bothaqueous and organic layers was analyzed by using HPLC with UV and fluorescencedetection. The aqueous layer (10 il) was directly injected into the HPLC and the organiclayer was analyzed after the solvent (CH2C1)was changed to acetonitrile as follows. Aportion (5 ml) of the organic layer was pipetted into a small glass sample vial and theCH21 was allowed to evaporate to dryness under a gentle N2 stream. The residue wasimmediately dissolved in 0.5 ml of acetonitrile. The conditions used for HPLC analysis ofcomponents were the same as those previously described in Section 2.2.1.Another set of experiments was carried out in order to monitor the amount of14C02 in the culture flask during the incubation. Round bottom flasks (250 ml) with twoside arms were used as culture flasks, so that a portion of the culture could be taken outfor analysis from one side arm and the same amount of replacement gas and liquid mediumcould be introduced from another side arm to balance the pressure inside the flask. Inaddition to the pyrene (60 p.g unlabeled and 200 nCi labeled) and enriched culture (1 ml)as described above, a volume of 100 ml of MS medium was added to each sterile roundbottom flask. At appropriate intervals during the incubation, samples composed of 5 ml53liquid and 10 ml head space gas were withdrawn from each flask, by using disposablesterile 20-ml Luer-Lok syringes fitted with appropriate needles. The sample wasimmediately injected through a serum stopper into a 25-mi bottle that contained 4 ml of 8M sulfuric acid. The14CO2released was flushed, trapped, and subsequently determined byusing the method described above. This sampling procedure removed approximately1/20th of the liquid and head space gas. It resulted in a slight vacuum in the culture flask,as the flask was sealed with serum rubber stoppers. To restore the pressure in the cultureflask, 5 ml of liquid medium and 10 ml of head space gas were transferred aseptically fromone of the control flasks and injected through a side arm into the culture flask. Theresulting vacuum in the control flask was balanced by the addition of 15 ml of air filteredthrough a sterile disposable bacterial air filter disc that was fitted on a Luer-Lok syringe.3.2.9. Acetylation and GC analysisThe acetylation method was adopted from that reported by Brooks et al. (212).Approximately 100 jig of standard cis-4,5-dihydroxy-4,5-dihydropyrene was treated withacetic anhydride (20 p1) and pyridine (10 jfl) and this mixture was placed in a warmwater bath at 60 OC for about 30 mm. Then the reagents were evaporated to drynessunder a gentle N2 stream. The residue was dissolved in toluene (approximately 100 p1)and analyzed by using both GC/FID and GC/MS as follows. An aliquot of this solutionwas injected to the GC injector at 260 OC and the components were separated by using aDB5 or a PTD 5 capillary column (30 m x 0.25 mm i.d.). The separation was carried outunder the following temperature program: an initial temperature 100 °C for 1 mm, thenramping at 5 °C/min to 300 °C, and finally holding at 300 °C for 15 mm.Three flasks of the cultures containing Pysed- 1 were extracted by using CH21,and this extract was evaporated to dryness. The residue was then acetylated and analyzedas described above.543.3. Results and Discussion3.3.1. Degradation of pyrene by enriched culturesPreliminary investigations into the degradation of pyrene were carried out byincubating 10 ml of mineral salt medium containing 10 pg of pyrene with 0.5 ml of a seawater culture (Pyw-2) enriched with pyrene. The concentration of pyrene in the culturewas monitored by using a scanning UV/visible absorption spectrophotometer. Figure 3.3shows three superimposed UV/visible absorption spectra obtained from the same set ofcultures that were sampled at incubation times 0, 72, and 108 hr. respectively. It is clearlydemonstrated that, as incubation time increases, the concentration of pyrene in the culturedecreases. Only approximately 30% of the pyrene originally added was present in theculture after 72 hr of incubation at ambient temperature. When the incubation time wasincreased to 108 hr, little pyrene remained in the culture. No loss of pyrene was observedin the sterile control, indicating that the disappearance of pyrene in the culture can beattributed to microbial degradation.Quantitative determination of the pyrene concentration in the sea water culturewas carried out by using HPLC with fluorescence and UV absorption detection. Threereplicate control and culture samples at each incubation time were analyzed. Thepercentages of pyrene in the culture relative to its initial concentration (1 .tg/m1) were42%, 5%, and 0, at incubation times 72, 120 and 170 hr, respectively; whereas the pyrenein the controls remained unchanged during the incubation. It is clear that as incubationtime progresses pyrene in the culture is degraded and that complete degradation of 10 igof pyrene is observed after 170 hr of incubation.It is important to point out that pyrene is the only organic compound added inthese experiments: the mineral salt medium used contains inorganic nutrients only. Thefact that pyrene is degraded in the present experiment suggests that the microbial550U,000RiU-’000o 200000RiU,0community can use pyrene as its sole organic substrate. There are only few suchexamples (100, lOOa, 101) of the microbial degradation of pyrene.0Cc.’D240 300 360 42000500 00Absorption Wavelength, nmFigure 3.3. Absorption spectra of a culture (Pyw-2) at incubation time(a) 0, (b) 72, and (c) 108 hr.56In addition to the sea water culture described above, five other cultures of marinesediments and sea water, that were enriched with pyrene or phenanthrene as the solesource of carbon and energy, were also used to carry out degradation studies. Thecultures containing 10 jig pyrene, 10 ml mineral salt medium and one of the inoculants(0.5 ml) were prepared and incubated. The concentration of pyrene in each flask wasdetermined by using HPLC with UV and fluorescence detection, and the percentage ofpyrene remaining in the culture, relative to the initial concentration, is summarized inTable 3.3. The results show that pyrene is rapidly degraded by using both the sedimentand sea water cultures enriched with either pyrene or phenanthrene. Approximately 31-56% of the total pyrene (10 jig) is degraded after 48 hr, and a complete degradation isachieved after 120 hr of incubation.Table 3.3 Degradation of pyrene by using enriched culturesIncubation time, hr. 0 48 112 120Culture Percentage of pyrene left, %Control 100 97±5 90±10 90±10Pysed-1 100 50±14 19±9 8Pysed-2 100 57±22 0.1 ndPyw-1 100 21±6 1±2 ndPyw-2 100 66±6 - ndPhensed-1 100 47±11 0.1 ndPhensed-2 100 91±4 - 5.1nd-not detectable (<1 nglml)- not evaluated57HPLC analysis of the culture solutions over time revealed the presence of a newcompound, presumably a metabolite, in addition to the pyrene. As an example, Figure3.4 shows HPLC traces from the control, Pyw-l, and Phensed-l after 48 hr ofincubation. A peak at retention time 3.8 mm, which is absent in the control, is observedin all the six cultures studied. This peak is due to a metabolite from the degradation ofpyrene.Figure 3.5 shows the results of a time series study of the relative concentration ofpyrene and metabolite in cultures from a culture incubated with Pysed-2. The relativeconcentrations were obtained by taking the ratio of the peak area of pyrene, or themetabolite, measured in the cultures at a given incubation time, to that of total pyrene inthe culture at incubation time 0 (a constant value). The metabolite is not fully identifiedat this point in time and hence its absolute concentration in the culture cannot bedetermined. Thus the relative concentration is a good measure. As shown in Figure 3.5,the concentration of pyrene decreases as the incubation time increases. This is consistentwith results shown in Table 3.3. The concentration of the metabolite increases during thefirst 75 hr of incubation and gradually decreases afterwards. A maximum concentrationof the metabolite in the culture is observed when pyrene is reduced to about 30% of itsinitial amount. As pyrene is further degraded, the metabolite is also used up, presumablyby the microorganisms present in the culture.58(a)(b)(c)I I I I I I0 2 4 6 8 10 12 14 16Retention Time, mmFigure 3.4. HPLC traces obtained from (a) culture Phensed-l, (b) culture Pyw-1,and (c) control after 48 hr of incubation.HPLC column: SupelcosilTM LC-PAH, 300 x 3.9 mm;mobile phase: 70% acetonitrile in water at a flow rate of 1.0 mI/mm.p - pyrene and m - presumed metabolitepmp59100I E200Incubation Time, hrFigure 15. Relative concentration of pyrene and a presumed metabolite measured at severalintervals during incubation of pyrene and an enriched sediment culture Pysed-2.(0 ) pyrene in control(• ) pyrene in the culture( ) the metabolite in the culture0 50 100 15060A similar experiment was carried out by using a culture, that originated from asea water sample, Pyw-2; more frequent sample monitoring was adopted in order tofurther investigate variation in the metabolite concentration. Samples from triplicatecultures were analyzed for pyrene and the metabolite at 12 hr intervals and relativeconcentrations of the two compounds are shown in Figure 3.6. A lag phase ofapproximately 45 hr is observed for the degradation of pyrene. Following this lag phasedegradation of pyrene takes place rapidly (a log phase). Degradation of pyrene to anundetectable level occurs after a total of 120 hr of incubation. The concentration of themetabolite increases during the log phase of pyrene degradation and reaches a maximumwhen pyrene is reduced by 40-70% compared to its initial amount. This maximumconcentration of metabolite remains for 25 hr of incubation time before it is graduallyreduced to undetectable level. The metabolite is probably further degraded (ormineralized) to smaller molecules as will be discussed in the next section. Nodegradation of pyrene nor its metabolite was observed in the sterile control. Generally,the maximum amount of the metabolite was observed in both the sea water culture andthe sediment culture when pyrene was degraded to less than 50%.The metabolite eluted from HPLC at the same retention time was also found inpyrene degradation studies involving other cultures. Table 3.4 shows the peak area of themetabolite from HPLC analysis of cultures at various incubation times. The metabolite isdetected in all these cultures and is absent in the control. The maximum production ofthis metabolite appears at different incubation times when different cultures are used asinoculant. This is likely because both the population of pyrene-degradingmicroorganisms and the degradation rate differ when these different cultures are used.61120c-)::400200Incubation Time, hrFigure 3.6. Relative concentration of pyrene and a metabolite measured during theincubation of pyrene and an enriched sea water culture Pyw-2.(0 ) pyrene in control(• ) pyrene in the culture(B ) the metabolite in the culture0 50 100 15062Table 3.4 Peak area of the metabolite (3.8 mm) produced in culturesIncubation time, hr. 0 48 112 120Culture Peak areaControl 0 0 0 0Pysed-1 0 13480 5808 0Pysed-2 0 11664 26918 13976Pyw-1 0 28726 NA 0Pyw-2 0 16489 NA 0Phensed-1 0 24226 19448 NAPhensed-2 0 0 NA 23314NA- not analyzed due to loss of sample3.3.2. Mineralization of pyreneIn order to investigate if the metabolite from pyrene degradation is furthermineralized to CO2 and H20, radioactive tracer studies involving 14C-labeled pyrenewere carried out. Mineralization of pyrene to produce CO2 and H20 was assessed bymeasuring the radioactivity of ‘4C02 produced from a culture containing a knownamount of(14C-4,5,9,10)-labeled pyrene. Both Pysed-1 and Pysed-1-Y were usedseparately as inoculants and duplicate cultures and controls were included in this study. Itwas found that approximately 65% or 55% of the radioactivity of(14C-4,5,9,l0)-pyreneadded into the cultures was mineralized to 14C02, 12 days after 14C-pyrene was63incubated with Pysed-1 or Pysed-1-Y cultures, respectively. No ‘4C02 was detected inthe controls, indicating that mineralization did not take place without microbial activity.Following the purge and trapping of‘4C02, the aqueous cultures were extractedwith methylene chloride. The radioactivity due to 14C in both the methylene chloridelayer and the water layer was measured. The recovery of the ‘4C added as pyrene into thecultures is 98%. The relative percentages of ‘4C in the two phases compared with thetotal 14C added into the culture are summarized in Table 3.5. The residual 1-4C in thePysed-1 and Pysed-1-Y cultures after 12 days of incubation is largely extracted into themethylene chloride phase.Table 3.5. Contents of 14C in organic and aqueous layersCulture 4CO2 (%) Residual 14C, (%) Pyrene (pg)Aqueous Organic (labeled + unlabeled)Control 0 0 98±2 62Pysed-l 65±6 7±2 25±5 0.7Pysed-1-y 55±6 8±3 35±5 3.0Extracts of the cultures and controls were further analyzed to quantitativelydetermine pyrene and its metabolites in the control and culture residues by using HPLCwith UV absorption and fluorescence detection. The results are also summarized in Table3.5. The total amount of pyrene in the control is 62 p.g. This is the amount of pyreneincluding both labeled and unlabeled pyrene initially added to each of the cultures andcontrols. However, only 0.7 ig of pyrene is left in the culture Pysed-1 and 3 ig in the64culture Pysed-1-Y after 12 days of incubation, accounting for 1% and 5%, respectively,of the total pyrene added initially. Therefore, the‘4C-containing components extractedinto the methylene chloride fraction is mainly from metabolites and cell carbon. A traceamount of the metabolite at the retention 3.8 mm described above was observed in bothcultures from HPLC traces. Thus other unidentified metabolic products containing ‘4Cmight be responsible for the balance of the radioactivity in the culture solutions.The results in Table 3.5 also show that similar amounts of 14C labeled pyrene aremineralized to give‘4C02 by the mixed cultures Pysed-1 and Pysed-1-Y. Pyrene is theonly source of carbon supplied to the culture in these studies. Thus the culture Pysed-1-Y, containing 2 strains as shown in Table 3.2, may be entirely responsible for theutilization of pyrene as sole source of carbon and energy in the mixed sediment culturePysed-l.The estimated mineralization of pyrene by the enriched sediment culture Pysed-1was studied over time. The percentages of pyrene mineralized at a sequence ofincubation times were measured and are shown in Figure 3.7. It is clear thatmineralization of pyrene to CO2 and H20 occurs quickly in the Pysed- 1 culture, up to45% of the pyrene is mineralized after 12 days of incubation. Further incubation to 38days results in 65% mineralization of the total pyrene added. A slower mineralization isobserved during the later phase of the incubation, probably because of a combination ofthe depletion of oxygen, the reduced supply of carbon and energy sources as pyrene andits metabolites are degraded, and the increased CO2 content in the closed incubationvessel. No‘4C02is observed from the sterile controls, confirming that the mineralizationof pyrene in the cultures is due to microbial activity. Again, pyrene is the sole source ofcarbon and energy in the mineralization study, no other source is provided.657050C—4.)10-101000Incubation Time, hrFigure 3.7. Estimated percentage of pyrene mineralized following the incubation of‘4C-labeled pyrene and an enriched sediment culture Pysed- 1.(0) in control and (• ) in the culture Pysed-10 200 400 600 800663.3.3. Identification of metabolitesThe identification of metabolites is essential for understanding microbialdegradation mechanisms and was therefore studied in detail. In the present investigations,metabolite identification was initially carried out by using (i) HPLC with variouscolumns and mobile phases, (ii) UV/visible spectra, and (iii) fluorescence excitation andemission characteristics. A number of studies (112, 195, 202, 213) have identifieddihydrodiol derivatives of PAHs as common metabolites, and degradation mechanismshave been proposed. Cerniglia’s group (101) has reported that cis-4,5-dthydroxy-4,5-dihydropyrene is one of the major metabolites from pyrene in bacterial cultures. Thus thiscompound was the primary target in present studies. An authentic standard of thisdihydrodiol was kindly supplied to us by Dr. Cerniglia.Figure 3.8 shows HPLC chromatograms obtained from (a) the metabolite from aculture sample, (b) the standard cis-4,5-dihydroxy-4,5-dihydropyrene, and (c) a 1:1mixture of (a) and (b). The HPLC column used (Supelcosil LC-PAH column (4.5 x 25mm, 5 pm)) is particularly efficient for the separation of mixed PAHs and their analogs.As shown in Figure 3.8, the same HPLC retention time (10.9 mm) for the metabolite andthe standard compound was obtained, when methanollwater (55/45) was used as mobilephase flowing at 1 mllmin. Under these conditions, pyrene was not eluted from thecolumn until 50 mm later. Changing the mobile phase to 55% acetonitrile in waterresulted in a shorter retention time (4.5 mm) for both the metabolite and cis-4,5-dihydroxy-4,5-dihydropyrene.67(a)(b)16 20 24 28 32Retention Time, mmFigure 3.8. HPLC/ fluorescence traces obtained from (a) the culture, (b) standard cis-4,5-dihydroxy-4,5-dihydropyrene, and (c) 1:1 mixture of (a) and (b).HPLC column: Supe1cosi1 LC-PAH, 300 x 3.9 mm;mobile phase: 55% methanol in water at 1.0 mI/mm., fluorescence excitationand emission wavelengths 260 nm and 370 nm, respectively.68When another reversed phase C18 column, ODS-2 (GL Scientific) was usedalong with methanol/water mixture (55/45) as mobile phase, both the metabolite in theculture and cis-4,5-dihydroxy-4,5-dihydropyrene eluted at the same retention time, 25.6mm. Analysis of a culture sample spiked with the standard (1:1 mixture) confirmed thatthe retention time of the metabolite matched that of cis-4,5-dihydroxy-4,5-dihydropyrene.A sample of trans-4,5-dihydrodiol pyrene was not available to us. However, the cis- andtrans- forms are easily separated by using HPLC conditions described above asdemonstrated by Heitkamp et at. (101). These results strongly suggest that the metaboliteis cis-4,5-dihydroxy-4,5-dihydropyrene.Further information on the metabolite was obtained by comparing UV/visibleabsorption and fluorescence spectra of the metabolite and the cis-4,5-dihydroxy-4,5-dthydropyrene standard. A UV/visible absorption spectrum was obtained by scanning astandard solution of cis-4,5-dihydroxy-4,5-dihydropyrene. The maximum absorptionpeaks are found at 220 and 257 nm with a wide shoulder around 286 nm, in goodagreement with the literature(l01). Attempts to obtain a similar spectrum of the culturecontaining the metabolite were not successful because only trace amount of themetabolite was present in the culture and the scanning UV/visible spectrophotometer isnot sensitive enough for the measurement. However a UV/visible absorption spectrum ofthe metabolite was obtained by using HPLC/UV. The peak intensity of the metabolitemeasured at each wavelength was plotted, giving a complete spectrum as shown inFigure 3.9. For comparison, a spectrum of cis-4,5-dihydroxy-4,5-dihydropyrene obtainedin the same way is also shown in Figure 3.9. It is clear that the maximum absorptionpeaks in the two spectra are identical.Fluorescence excitation and emission spectra of the metabolite and cis-4,5-dihydroxy-4,5-dthydropyrene were similarly obtained and are shown in Figure 3.10.Again the fluorescence characteristics of the metabolite and standard cis-4,5-dihydroxy-4,5-dihydropyrene are identical.CCciC>==69543210200 220 240 260 280 300 320 340Absorption Wavelength, nmFigure 3.9. Absorption spectra of (• ) a metabolite in cultures and (0) standard cis-4,5-dihydroxy-4,5-diliydropyrene obtained by using HPLC/UV.See Figure 3.4 for HPLC conditions.704..—3.190 240 290 340 390 440Excitation EmissionWavelength, nmFigure 3.10. Fluorescence excitation and emission spectra of (• ) metabolite in culturesand (0) standard cis-4,5-dihydroxy-4,5-dihydropyrene obtained by usingHPLC/FluorescenceSee Figure 3.4 for HPLC conditions.At excitation wavelength 252 nm for obtaining emission spectra and atemission wavelength 370 nm for obtaining excitation spectra71Based on the identical HPLC retention time and absorption and fluorescencespectra between the metabolite and the standard, it is believed that cis-4,5-dihydroxy-4,5-dihydropyrene is the major metabolite in the present study of pyrene degradation. Inorder to further confirm the identification of this metabolite, GCIMS was used to obtainthe mass spectrum of the metabolite. cis-4,5-dihydroxy-4,5-dihydropyrene wasdecomposed to pyrene, pyrenol and other compounds in the GC injector (260 oc). Tosolve this problem, cis-4,5-dihydroxy-4,5-dihydropyrene was acetylated by treatmentwith acetic anhydride and pyridine. The product from the acetylation of the standard cis4,5-dihydroxy-4,5-dihydropyrene was analyzed by using both GC/FID and GC/MS. Onlyone peak was observed from both the GC/FID and GCIMS analysis. This suggests thatthe acetylation of the standard cis-4,5-dihydroxy-4,5-dihydropyrene was complete. AGCIMS trace of the acetylation product from the standard cis-4,5-dihydroxy-4,5-dihydropyrene is shown in Figure 3.11(a) and the mass spectrum is shown in Figure3.11(b). The M peak at m/z 320 is due to the diacetate of the cis-4,5-dihydroxy-4,5-dihydropyrene. The other major fragment ions are assigned as follows: 260, loss ofCH3OOH from the M; 218, loss ofCH2O from mlz 260; and 189, further loss of Hand CO. Thus the acetylation product from the standard is identified as diacetate of cis4,5-dihydroxy-4,5-dihydropyrene. When the same acetylation treatment was applied tothe culture extract, a peak at the identical retention time to that of the standard was alsodetected by using both GC/FID and GC/MS. Figure 3.12 shows the chromatograms ofthe standard (a) and the culture sample (b), obtained by using GC/FID, and (c) the massspectrum of the metabolite in the culture, obtained by using GC/MS. The results inFigures 3.11 and 3.12 show that GC retention time and the mass spectrum of themetabolite after acetylation are identical to those of the standard, confirming that themetabolite from the culture is cis-4,5-dihydroxy-4,5-dihydropyrene.90 00 70 60 50-1126/40-30 20 10 o10:00—20:0030:0040:0050:00218.090 80-70 60 50-40-30189.00319.920-259.91:4.Oe8.__1.050234.91°•050100150___200250300350400m/zt.JFigure3.11.GCIMStrace(a)andmassspectrum(b)oftheacetylationproduct ofcis-4,5-dihydroxy-4,5-dihydropyrene730.O20.02C (a)o.oie 23.0750.010.O10 oiL - -0.Ol81D1(b)23.075O.Oi2001 10 10 20218 (c)-70-40-260 3201o-o_ .. -.,.0 50 100 150 300 350Figure 3.12. The GC/FID traces of the acetylation products of (a) standard ciS4,5-dihydroxy-4,5-dihydropyrcne (b) culture extract, and (c) mass spectrum of (b) by GCIMs.74Cis-4,5-dihydroxy-4,5-dihydropyrene is an initial ring oxidation product frompyrene. It has been proposed (94, 102, 105, 106) that bacterial degradation of PAils isinitiated by a dioxygenase. Two atoms of molecular oxygen are incorporated into anaromatic ring of the PAil, resulting in a cis-dihydrodiol derivative. Thus cis-dihydrodiolderivatives of naphthalene (112, 195, 213), phenanthrene (119, 202), anthracene (119,202), and pyrene (101) have been found to be the initial ring oxidation products duringthe degradation of these PAHs. In the co-metabolism of pyrene by Mycobacterium sp.with co-existing of organic nutrients (101), however, both dioxygenase andmonooxygenase enzymes were involved, resulting in both cis- and trans-dihydrodiolpyrene as initial ring oxidation products. In the present study, only cis-4,5-dihydroxy-4,5-dihydropyrene is detected in all the cultures, but trans-4,5-dihydroxy-4,5-dihdropyrene isnot produced. This suggests that a dioxygenase is responsible for the metabolism, withpyrene being the sole source of carbon and energy; a possible mechanism involves initialoxidation of the double bond at the 4,5 positions. In the only other report (100) of thebacterial utilization of pyrene as sole carbon source by Rhodococus sp. UW1, the majormetabolite isolated was a ring fission product. No initial ring oxidation product wasobserved. Based on the ring fission products, the pathway for the initial oxidation ofpyrene by this species could involve either the 4,5 or 1,2 positions. The present studysuggests that the pathway of bacterial utilization of pyrene as sole source of carbon,follows initial attack on the 4,5 position to form the cis-4,5-dihydroxy-4,5-dihydropyrene,and that a dioxygenase is responsible for the initial ring oxidation.It is interesting to note that the fluorescence properties of the metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene, are different from those of pyrene but are similar to thoseof phenanthrene. As shown in Figure 3.13, a very small peak from the metabolite isobserved when the optimum excitation and emission wavelengths for pyrenedetermination were chosen, i.e. 238 and 338 nm (158), respectively. However, theintensity of the metabolite peak is significantly enhanced when the optimum excitation75(252 nm) and emission (370 nm) wavelengths for phenanthrene determination are used.These results suggests that the fluorescence excitation and emission wavelengths shouldbe carefully selected for the determination of PAH degradation metabolites. Furtherstudies on the fluorescence characteristics of a variety of possible metabolites shouldprovide useful information in this regard.Figure 3.13. Fluorescence intensity of a culture supernatant at(a) Excitation 238 nm and Emission 338 nm, and(b) Excitation 252 nm and Emission 370 nmSee Figure 3.4 for HPLC conditions.(a)(b)‘a)‘a)C.)‘a)CD‘a)1-CI I I I0 2 4 6 8 10 12 14 16Retention Time, mm76In addition to cis-4,5-dihydroxy-4,5-dihydropyrene, another metabolite isobserved in some cultures. Figure 3.14 shows chromatograms obtained from sterilecontrols and cultures incubated for 2 days (a) and 6 days (b) after being inoculated with asea water culture strain, Pyw- i-y. Cis-4,5-dihydroxy-4,5-dihydropyrene has a retentiontime of 3.8 mm, but a new peak at retention time 6.0 mm was observed in both culturesamples. These peaks are not observed in any of the sterile controls (Figure 3.14 c).Similarly, when another culture, Pyw-2 (10 ml), was inoculated into 400 ml of mineralsalt medium containing 500 ig of pyrene, the peak at 6.0 mm. was also clearly observedas shown in Figure 3. 15a.This metabolite was initially thought to be pyrenol because pyrene has beenreported to be a metabolite from the degradation of pyrene by using MycobacteriumPRY-i (101). The metabolite eluted at the same retention time as pyrenol from theSupe1cosil LC-PAH column when 70% acetonitrile in water was used as mobile phase(Figure 3.15 a ,b, and c). However, it was soon established that the fluorescenceproperties of this metabolite are distinctly different from those of pyrenol. Whilestandard pyrenol was detected (Figure 3.15 e) by using the fluorescence detector atexcitation wavelength 260 nm and emission wavelength 370 nm, the metabolite was notobserved under these conditions (Figure 3.15 d). The fluorescence spectrum of themetabolite is shown in Figure 3.16. The UV spectrum for the metabolite (Figure 3.17)also shows different maximum absorption wavelengths from those of pyrenol. Therefore,it is clear that this metabolite is not pyrenol. In fact further optimization of HPLCconditions (55% acetonitrile in water as mobile phase) results in baseline separation ofthe metabolite and pyrenol. Although the present results do not provide a conclusiveidentification of the metabolite, the spectral and chromatographic characteristics of themetabolite will be useful in further studies of the metabolites from PAH degradation.77(a)It II(c)0 2 4 6 8 10 12 14 16Retention Time, mmFigure 3. 14. HPLC/UV traces of (a) Pyw-1-y at 2 days, (b) Pyw-1-y at 6 days,and (c) control at 6 days, following the incubationSee Figure 3.4 for HPLC conditions.p. pyrene; I and II - metabolites.78II’0 2 4 6 8 10 12 14 16Retention Time, mmFigure 3. 15. HPLC traces obtained using UV detection at 254 nm (a, b, and c) andfluorescence detection at excitation 260 nm and emission 370 nm (d, e, and f).a, d - culture sample; b, e - standard pyrenol (0.13 .tgIm1); andc, f -1:1 mixture of the culture sample and the standard pyrenol solution.(a) (d)(b) (e)(c) (f)I I0 24 6 810121416Retention Time, mm79—l-4.-(-.t•)Q0cjDCzFigure 3. 16. Fluorescence spectrum obtained from a metabolite in the culture (retentiontime 6.4 mm).At excitation of 250 nm to obtain the emission spectrum andat emission of 420 nm to obtain the excitation spectrum.210200Excitation300 400 500EmissionWavelength, nm80864.20- I • I • •200 250 300 350 400Absorption Wavelength, nmFigure 3.17. Comparison of UV absorption spectra obtained from ( A ) pyrenol standard;(0 ) cis-4,5-dihydroxy-4,5-dihydropyrene standard; and (•) a metabolite inthe culture.81Other possible metabolites were also observed. After the pyrene was completelydegraded, a 2 ml aliquot of the supernatant of the culture Pyw-2 was subjected to apreparative C18 column for separation. The chromatogram in Figure 3.18 was obtainedwhen 50% acetonitrile in water was used as mobile phase at flow rate 6.8 mllmin. Figure3.18 shows four major peaks at retention time 4.8, 6.0, 7.8, 10.0 mm., respectively. Fourfractions were collected from the four peaks. These fractions were condensed to drynesson rotary evaporator at a temperature of 40 °C. They were then analyzed by usingdesorption chemical ionization (CI) mass spectrometry. Fraction I was also analyzed byusing electron impact ionization (El) mass spectrometry. As an example, the CI and Elmass spectra of fraction I are shown in Figure 3.19. Both CI and El spectra are similar toeach other, except that the El spectrum has more fragment ions at lower miz ratio.Fraction I has M 242 and major fragment ions at mlz 186 and 142. The M 242corresponds to the formula C15H1403, the fragment ions 186 to loss of CH2=CHCHO,and 142 to loss of CH2=CHCHO and C02. The m/z 142 and 128 suggest‘CHandrespectively. Thus the fraction I is tentatively identified asCHH HOCHçOOH820CIIIIIv1’I I I I I0 4 8 12 16 20Retention Time, mmFigure 3.18. HPLC/UV trace obtained from an original culture Phensed-2.On a preparative column: Waters, C18, 100 x 25 mm.Mobile phase: 55% acetonitrile in water, 6.8 mllmin.100%-II-100%161250%50%(a)18,1-0%50100150200250m/z100142(b)10070 60 501404130 20j423269911281850liiijIIII406080100120140160180200220240Figure3.19. MassspectraoffractionIcollectedfrompreparativecolumn,using(a)chemicalionizationand(b)electronionization84The other fractions are not identified. The peaks observed above indicate thatmany metabolites are present in the cultures. This is consistent with the 14C tracer studiesthat the major 14C activity left in the culture residues was due to the metabolites and onlyminor 14C-labeled pyrene was detected. These results suggest that the identification ofother metabolites warrant further study.3.3.4. Degradation of pyrene by original culturesThe studies described above involved the use of subcultures in which certainmicroorganisms were selectively enriched from sea water and sediments by serialtransfer into mineral salt medium and pyrene. These enriched cultures show the ability toutilize pyrene as sole carbon source. Further studies were carried out by using theoriginal sea water and sediment cultures.Sixteen cultures were generated by inoculating sea water and sediment into (i)mineral salt medium, (ii) mineral salt medium and phenanthrene, (iii) mineral saltmedium and pyrene, and (iv) mineral salt medium and benzoanthracene. These cultureswere kept on a shaker for six months before further investigation. Each culture was usedas inoculant in the following experiments to study the degradation of pyrene as follows.Each of the above cultures (0.5 ml) was inoculated into 10 ml of mineral salt mediumcontaining 10 jig of pyrene, and the concentration of pyrene in the culture afterincubation was determined and compared with the initial amount added. The averagepercentage of pyrene remaining in the cultures relative to the initial concentration aresummarized in Table 3.6. Each mean value is obtained from the analysis of triplicatesamples by using HPLC with both UV and fluorescence detection. Results in Table 3.6show that pyrene is degraded by eight cultures after 9 days of incubation. As theincubation proceeds to 15 days, all the sixteen cultures studied show the ability to degrade85pyrene; pyrene being the only carbon source supplied to the culture. Among thesecultures, four of them directly originated from sea water and sediment without adding anyPAH. Results in Table 3.6 demonstrate that these mixed cultures also remove pyrene.Previous studies by others (123, 127, 129, 191, 210) have also suggested thatmicroorganisms present in PAH-contaminated environment have higher ability to degradePAHs.The degradation of pyrene by the sixteen original cultures was further investigatedby examining simultaneously both the decrease of pyrene concentration and theappearance of metabolites. The HPLC peak intensities of the metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene, in cultures incubated for various time periods are listed inTable 3.7. The expected metabolite is found in almost all of the cultures. The onlyexception is the culture, Water-i no PAH, where the metabolite is not detected, probablybecause its concentration is below the detection limit (5 ng/ml). The maximum amount ofthe metabolite in different cultures appears at different incubation times, suggesting thatthere were different populations of microorganisms and different degradation ratesassociated with the various cultures.86Table 3.6. Percentage of pyrene left after pyrene incubated with each ofthe sixteen original culturesCulture Incubation time, day3 6 9 12Killed cell control, sediment 90±9 93±2 87±2 86±6Killed cell control, sea water 97±6 - 89±3 -Reference sediment-i 83±5 84±6 83±3 ndReference sediment-2 91±6 68±10 8 ndReference water-i 86±11 90±5 89±1 59±6Reference water-2 84±7 nd nd ndPhenanthrene sediment-i 82±1 - ndPhenanthrene sediment-2 56 - nd -Phenanthrene water-i 86±2 85±6 91±1 60±3Phenanthrene water-2 89±1- 5±4Pyrene sediment-i 86±8- 1±1 ndPyrene sediment-2 47±1 - ndPyrene water-i 85±7 - 72±6 ndPyrene water-2 73±4 - ndBenzoanthracene sediment-i 88±5- 76±13 58±7Benzoanthracene Sediment-2 97±2 - 53±20 ndBenzoanthracene water-i 81±5 82±3 80±6 1±2Benzoanthracene water-2 101±14 - 78 58±7nd-not detectable- not evaluated87Table 3.7 Peak area (x104)of the metabolite produced by the sixteenoriginal culturesIncubation time, dayCulture 3 6 9 12Killed cell control, sediment nd nd nd ndKilled cell control, sea water nd - ndReference sediment-i nd nd- 0.78±0.04Reference sediment-2 0 1.0±0.4 1.21 -Reference water-i nd nd nd ndReference water-2 nd 2.7±0.4 2.7±0.4 0.7±0.1Phenanthrene sediment-i nd-5.2±5.5Phenanthrene sediment-2 1.6 0.58Phenanthrene water-i nd nd 0.56 0.7±0.2Phenanthrene water-2 nd- 6.6±3.7 ndPyrene sediment-i nd 2.2±0.6 0.7Pyrene sediment-2 1.62±0.03 nd nd ndPyrene water-i nd i.5±0.4 nd -Pyrene water-2 2.2- 2.04 ndBenzoanthracene sediment-i nd nd nd 0.3Benzoanthracene Sediment-2 nd- i.i3±0.02 2.0±0.1Benzoanthracene water-i nd nd 0.96 2.1±0.1Benzoanthracene water-2 nd nd nd 0.3nd-not detectable883.3.5. Degradation of pyrene by isolated strainsAs described in Section 3.2.5, isolation was carried out from four mixed cultures,giving eleven strains. Characteristics of these pure strains are summarized in Table 3.2.Eight of these isolated strains were used as inoculant for further degradation studies, inorder to investigate their ability to degrade pyrene.Eight cultures including Pysed-l-y, Pysed-l-ly, Pysed-2-y, Pysed-2-ly, Pyw-l-y,Pyw-1-ly, Pyw-2-y, Pyw-2-ly, Phensed-2-y, and Phensed-2-ly, isolated on the marinebroth agar plates, were grown separately on slants and transferred back to freshlyprepared liquid enrichment medium containing mineral salt and pyrene. After about threemonths of incubation, these eight cultures were studied for their ability to utilize pyrene assole carbon source. Each of the eight cultures (0.5 ml) was inoculated separately to themineral salt medium (10 ml) containing 10 jig pyrene, and the concentration of pyrene inthe cultures was monitored. Significant degradation of pyrene was observed only whenPyw-1-y and Pysed-l-y were used as inoculants. Complete degradation of pyrene to anundetectable level was obtained 71 hr and 102 hr after 10 jig of pyrene was incubated withPysed-1-y and Pyw-1-y, respectively. An example of the scanning UV spectrum fromPysed-1-y cultures is shown in Figure 3.20. These spectra shown in Figure 3.20 are similarto those in Figure 3.3 where enriched mixed cultures were used; degradation of pyrene isclearly illustrated.Similar results were also obtained when Pyw-1-y was used as inoculant. However,no degradation of pyrene was observed when the other six isolated strains were used asinoculants for pyrene degradation studies.Further isolation and purification on CGY media confirmed that the above sixstrains, which did not degrade pyrene, were all pure colonies. However, each of the twoisolates (Pysed-1-y and Pyw-1-y) which utilized pyrene as sole carbon source contained atleast two strains with distinct colony morphologies. These colonies are very similar and0000CC,)<0000000r\)0000C8900o 200C240 300 360 42000500 0CAbsorption Wavelength, nmFigure 3. 20. Scanning UV absorption spectra obtained from a culture containing isolatedstrain, Pysed-1-y.(a) pyrene in culture at incubation time 0(b) 72 hr after incubation.90Further isolation and purification on CGY media confirmed that the above sixstrains, which did not degrade pyrene, were all pure cultures. However, each of the twocultures (Pysed- l-y and Pyw- l-y) which utilized pyrene as sole organic substrate containedat least two strains with distinct colony morphologies. These strains are very similar anddiffer only in size and response to one out of the 95 biochemical reactions in the biologtests as described in Table 3.2.Purified strains, Pyw-l-yl (larger size) and Pyw-l-ys (smaller size), were grownseparately in mineral salt medium containing pyrene particles for seven weeks, and thesetwo pure cultures were used as inoculants to further study the degradation of pyrene.Surprisingly, no degradation of pyrene was observed during five weeks of incubation.Similar results were obtained when both Pyw- 1 -yl (larger size) and Pyw- 1 -ys (smaller size)were inoculated into the mineral salt medium containing pyrene: no degradation wasobserved. It seams that the pure strains lost their ability to degrade pyrene as sole carbonsource, possibly because they were grown on organic rich media prior to the isolation andpurification. The microorganisms may have been accustomed to using these carbon sourcesin the media which are easier to metabolize, thereby losing the ability to metabolize pyreneas sole carbon source. Another possibility is that several types of microorganisms may beneeded in the culture in order to develop capacity for utilizing pyrene as sole carbon sourceor, a third strain capable of degrading pyrene was not isolated from the mixed culture.Similarly, although Pysed-2 degraded pyrene rapidly by using pyrene as sole carbonsource, its pure isolates (Pysed-2-y and Pysed-2-ly) did not show any ability to degradepyrene. When these two isolates were combined together in the cultures, degradation ofpyrene was not observed either. These results are consistent with those described above andpresumably the same explanations.The isolation of pure cultures capable of using pyrene as sole sources of carbon andenergy warrants further attempts, since only one pure strain Rhodococcus sp. UW1 (100)has been reported to be able to utilize pyrene as sole sources of carbon and energy.91within 16 days of incubation of 10 p.g pyrene, in 10 ml mineral salt medium, and an aliquotof the preserved culture. The HPLC analysis of the control and culture samples after 16days of incubation shows a metabolite at retention time 3.8 mm in addition to thedisappearance of pyrene (Figure 3.21). This metabolite is the same compound previouslyidentified as cis-4,5-dihydroxy-4,5-dihydropyrene. No metabolite nor significant loss ofpyrene was observed in the controls. These resulis suggest that the preservation ofcultures does not alter the capability of the microorganisms to degrade pyrene and that thismethod of preservation can be used for future collection of PAH-degrading marinemicroorganisms.The preserved cultures were grown on half strength marine broth for a weekbefore preservation. The fact that they did not lose the pyrene-degrading ability suggeststhat half strength marine broth is suitable for the isolation of PAH-degradingmicroorganisms. Previous results on the rapid degradation of pyrene by using purifiedculture strains, Pysed-2-y and Pyw-l-y, also support this conclusion. These strains wereisolated on half strength marine broth over 6 weeks after serial transfers on the agar plates.On the other hand, the CGY may be not suitable, as strains purified on this medium losttheir ability of degrading pyrene, as discussed above.92pI I I I I I I02468101214Retention Time, mmJ16(a)(b)Figure 3. 21 .HPLC/LJV traces obtained from a regenerated culture Pysed-2 that had beenpreserved for seven months.(a) pyrene in culture at incubation time 0(b) 16 days after incubation; metablite (m) observed.See Figure 3.4 for HPLC conditions.m93Chapter 4. Mineralization of Pyrene by IndigenousMicroorganisms in Sediment Samples4.1. IntroductionThe results described in the previous chapters show that mixed cultures enrichedfrom the Kitimat samples are capable of degrading phenanthrene and pyrene. Themicrobial degradation of pyrene was further confirmed by the production of metabolitesand 14C02 from 14C labeled pyrene. It now remains to establish if the microorganismsare able to carry out these degradations in the natural environment. It is very important toobtain this information in order to understand the fate of PAHs in a particularenvironment of study.Although hydrocarbon-degrading microorganisms are ubiquitous in mostecosystems (214, 215), it is usually very difficult to prove that degradation and/ormineralization actually occurs in the field, because of the uncontrolled conditions in thenatural environment (216). It is difficult to distinguish between biotic and abioticprocesses. However, laboratory assays can provide definitive evidence for microbialdegradation and sterilized samples can be used to determine abiotic losses. Thuscontributions from microbial degradation can be differentiated from abiotic loss (216).Furthermore, a mass balance can be obtained by performing the assay in a sealed vessel.Some studies have lead to a better understanding of many aspects of the biodegradationof organic compounds including genetic, physiological, kinetic and ecological (94, 106,134, 214, 217-222). Results obtained from laboratory studies have also been applied to insitu bioremediation of gasoline-contaminated aquifers and oil-contaminated soils (102,104, 214, 215, 223-229).A number of approaches can be taken to obtain evidence for the biodegradation inthe field, in situ biodegradation (216). These include (i) quantitative determination of the94contaminant of interest in samples collected at different times to show a decrease in itsconcentration over time; (ii) laboratory based microbial degradation studies underconditions that mimic the environment to show the potential of biodegradation in thefield; and (iii) searching for a particular metabolite of biodegradation in samplescollected from the field.It is very difficult to demonstrate microbial degradation in Kitiamt Arm bymonitoring the loss of PARs in the samples, because PAHs are being continuously addedto the environment from the nearby aluminum smelting plant. Without knowing the valueand nature of the PAH input, it is impossible to estimate any loss of PARs. Furthermore,weathering mass transport of sediment and water, and other abiotic processessimultaneously occur and contribute to changes in the concentrations of PAHs insamples. For these reasons, two other approaches, laboratory microbial degradation andthe determination of a typical metabolite in the sample, were chosen, to evaluate thepossibility of microbial degradation in the field of Kitimat Arm.A simple method commonly used to determine microbial activity towards PAHsin sediments is to study the mineralization of 14C-labeled PAN inoculated into thesediment collected from the field. The production of 14C02 is measured and theefficiency of the mineralization of a specific PAH by indigenous microorganisms ismeasured by the ratio of the activity from 14C02produced to that of the total 14C added(230-233). This method does not alter the integrity of the sediment since only trace levelsof the 14C-labeled PAH is added. Therefore, this approach was adopted in the presentstudy and the mineralization of 14C-pyrene by indigenous microorganisms in sedimentswas studied. The 14C-pyrene was only labelled at positions 4, 5, 9, and 10, so onlypotential mineralization was demonstrated, and total mineralization was indirectlyestimated.Another approach was also used to support the findings of biodegradation in thefield. cis-4,5-dihydroxy-4,5-dihydropyrene has been identified as a metabolite of the95microbial degradation of pyrene as discussed in Chapter 3. Further mineralization of themetabolite was observed only after pyrene was degraded to low level. Consequently, ifthe microbial degradation that occurs in the environment is similar to that in laboratorystudies, cis-4,5-dihydroxy-4,5-dihydropyrene is expected to be present in the samplesfrom Kitimat Arm, where there is a continuous input of pyrene. For this reason, effortwas made to determine cis-4,5-dihydroxy-4,5-dihydropyrene in samples from KitimatArm, and the finding of this metabolite in the environment would provide a strongevidence for the presence of in situ biodegradation of pyrene in Kitimat Arm.In order to better understand biological degradation processes in the environment,a number of factors reported to influence the degradation of PAHs (102, 106, 120, 133,139, 233, 236) were investigated with respect to the degradation of pyrene. These factorsinclude temperature, salinity (NaCl content), medium composition, pyrene content, andmicrobial population.4.2. Experimental4.2.1. Sampling locations and characteristics of samplesThe present study focused on the northern part of Kitimat Arm, as shown inFigure 1.5 (Chapter 1) and Figure 4.1. All the samples used in this chapter were obtainedfrom the locations shown in Figure 4.1. CD-i, CD-2, CD-3, and CD-6 are close to theshore, CD-4, CD-5, K5a, and KA7 are further away from the shore and away from theMean Smelter. C-sed and STN2 are also located near CD-i. CD-2 is located near upperpart of the Aican Dock, and CD-3 is near the D lagoon discharge, between Aican Dockand Hospital Beach. The area around CD-3 had been previoously dredged. Thus samplesfrom CD-3 may be affected. The six CD samples (CD-i to CD-6) were obtained inNovember, 1993. Surface sediments were collected from the top 5 cm by using stainlesssteel grab. Samples were placed in sterile glass jars and stored at 4 °C for a month before96they were used. The C-sed samples were obtained in June of both 1992 and 1993. Thesamples collected in 1992 were used to generate cultures that were used for the studies ofphenanthrene and pyrene degradation, and a year later a sample was obtained from thesimilar location to investigate the presence of cis-4,5-dihydroxy-4,5-dihydropyrene.STN2, K5a, and KA7 samples were collected in May 1991. They were freeze-dried andstored at 4 °C, before they were extracted for the determination of cis-4,5-dihydroxy-4,5-dihydropyrene.All sediment samples were brown and sandy except for samples obtained fromnear Yacht Club CD-i, STN2, and C-sed which were dark muddy sediments and had adistinct sulfide smell, suggesting that this area was under anaerobic condition. Theconcentrations of sixteen PAHs in these samples have been determined in our laboratory(66) and summarized in Table 4.1.Table 4.1. The concentrations (j.ig/g dry weight) of the sixteen PAHs in the sedimentsSample CD-i CD-2 CD-3 CD-5 CD-6Naphthalene 0.3 0.5 0.1 nd 0.1Acenaphthylene nd nd nd nd ndAcenaphthene 2.1 1.8 0.1 .01 0.3Fluorene 1.6 1.0 nd 0.1 0.2Phenanthrene 17.3 9.8 0.3 0.6 1.7Anthracene 3.6 2.2 nd 0.1 0.4Fluoranthene 37.5 18.4 0.6 1.2 3.2Pyrene 34.4 18.1 0.6 1.2 3.0Benz(a)anthracene 34.4 13.0 0.6 1.0 3.2Chrysene 25.4 11.3 0.5 0.9 2.4Benzo(k)fluoranthene 65.9 24.9 0.9 1.7 5.6Benzo(b)fluoranthene 25.6 9.3 0.5 1.2 3.4Benz(a)pyrene 56.4 20.4 0.6 1.0 4.2Dibenz(a,h)anthracene 25.0 5.7 0.4 0.4 2.0Indeno(i,2,3-cd)pyrene 92.8 23.3 0.6 1.4 6.8Benzo(g,h,i)perylene 105.0 21.7 0.9 1.4 7.5Total 527.8 181.5 6.7 12.4 43.997KA7NSS‘B’ALCAN : Lagoons’Ic-see STN2CD-iMINE7TEBAYCD-5KITIMATCD-6 ARMFigure 4.1. A map showing the location of sampling sites.984.2.2. Mineralization of pyrene by indigenous microorganisms in sedimentsA portion of each sediment sample (CD-i to CD-6) was mixed with the surfacewater in the grab sampler to form a slurry: six of such slurry samples were freeze-driedto determine the contents of water and sediment, and it was found that the approximateratio is 30% sediment (dry weight) and 70% water by weight.14C-4,5,9,iO-pyrene (20nCi) was added to each slurry. An additional 100 pg of unlabeled pyrene was added toCD-3. The culture flasks were sealed with serum stoppers immediately after the additionof 14C-pyrene. The samples were well mixed and placed on a shaker (136 rpm). Thetemperature during the incubation varied between 8 to 15 °C according to the ambientconditions. After 23 days of incubation, 10 ml of 8 M sulfuric acid was added to eachflask. The flasks were gently shaken for 5 mm to ensure that all the 14C02 in the culturewas released into the head space. An aliquot of 10 ml of the head space gas waswithdrawn by using a gas tight syringe and immediately injected into a liquidscintillation (LS) vial containing a mixture of liquid scintillation cocktail (3 ml),ethanolamine (0.67 ml), and ethylene glycol monomethyl ether (0.33 ml). Another vialcontaining the same mixture was also connected to the first vial, as shown in Figure 3.2,to ensure complete trapping of the 14C02, although it was found that this wasunnecessary as discussed in Section 3.2.8. No sediment was taken for the measurementsince 14C02 was completely released to the head space upon the addition of acid.Triplicate samples of head space gas were taken from each culture for 14C02measurement. The amount of 14C02 produced was obtained from the threemeasurements and the known total volume of the head space. The mineralizationefficiency was determined by the ratio of the radioactivity of the 14C02measured fromeach sediment relative to that of the 14C-labeled pyrene initially added. Duplicatecontrols for CD-i, CD-3, and CD-6 were prepared by autoclaving the sediments beforethe addition of 14C-pyrene. The CD-3 controls were made up to contain an additional100 tg of un-labeled pyrene. All the controls were incubated under the same conditions99as those for the cultures. The same procedures were used for trapping and measuring14C02. Only background levels of radioactivity was measured from the controls. Thusthere was no abiotic production of 14C02during the incubation. All the 14C02producedin the samples can be attributed to microbial activity.In order to study the enhancement of mineralization of pyrene in the sedimentsamples, 1 ml of the enriched culture Pysed-1 was added to the samples CD-i, CD-3, andCD-6. The other procedures including preparation, incubation and determination ofwere the same as described above. Similarly, parallel cultures were autoclavedand used as killed cell controls.4.2.3. Determination of dihydrodiol pyreneAll sediment samples were freeze-dried. A portion of 50 g of dried sedimentSTN2 was shaken overnight in 200 ml acetonitrile. The supernatant acetonitrile extractwas decanted and filtered through a Wattman 4 filter paper to remove the particles. Thefiltrate was concentrated to approximately 16 ml on a rotary evaporator at 35 °C, andthen further condensed to 5 ml by evaporation under N2 stream. The extract was filteredthrough a 0.45 p.m membrane filter, and an aliquot of 10 p.1 filtrate analyzed by HPLC. Apeak at retention time 10.9 mm corresponding to cis-4,5-dihydroxy-4,5-dihydropyrenewas observed, when 55% MeOH in water was used as mobile phase at a flow of 1.0mi/mm. The extract also contained a great number of other components.In order to separate the peak at 10.9 mm from other late-eluting components andto reduce the elution time, the extract (1 ml) was first injected to a preparative column(Waters RCM 100 mm x 25 mm reversed phase C18); 55% MeOH in water was used asmobile phase at 13.8 mI/mm. A fraction eluted between 3 to 14 mm was collected,because a standard cis-4,5-dihydrodiol pyrene was eluted in this range of retention time.The fractions collected from several injections were pooled together, concentrated to 0.5ml, and an aliquot (10 p.1) of the concentrated fraction was then injected to an analytical100column (SupelcosilTh4LC-PAH) and eluted by using MeOHIH2O(55/45) at flow rate of1.0 mLlmin. The component which eluted at 10.9 mm, corresponding to cis-4,5-dihydroxy-4,5-dihydropyrene, was collected, pooled, and concentrated to 0.2 ml forfurther characterization purpose. When this purified fraction was analyzed on anotheranalytical column (ODS-2 GL Scientific), by using the same mobile phase, a peak at 25.6mm was detected, which matched the cis-4,5-dihydrodiol pyrene standard. This purifiedfraction (10.9 mm) was also used to study the UV absorption and fluorescence propertiesof the metabolite by using HPLC/UV/fluorescence detection. The absorption wavelengthwas manually changed on the detector. An absorption spectrum (200 to 320 nm) of themetabolite was obtained by using HPLC/UV; a Supelcosil LC-PAH column was used forseparation and 70% acetonitrile was used as mobile phase. The metabolite in the extractwas eluted out at retention time 3.8 mm under these conditions, and matched theretention time of the cis-4,5-dihydroxy-4,5-dihydropyrene standard.The fraction collected from the preparative column (Waters RCM) was acetylatedwith treatment of acetic anhydride and pyridine. The acetylation products were analyzedby using GCIFID and GC/MS. The procedures for the acetylation and determination ofthe products used here were the same as those described in Section 3.2.9. The sametreatment was also applied to the whole acetonitrile extract without pre-separation.However the expected compound co-eluted with other major components in the sample,so no mass spectrum of the metabolite was obtained from GC/MS. Therefore, it isnecessary to preseparate the metabolite from the other components in the acetonitrmleextract before GC/MS analysis in order to obtain a good mass spectrum of themetabolite.Other sediment samples were extracted with acetonitrile and these extracts weredirectly analyzed by using HPLC (Supelcosil LC-PAH) to screen the presence of themetabolite and to estimate its concentration. The mobile phase for the initial 15 mm was55% acetonitrile in water, then ramped to 100% acetonitrile in 20 mm. The mobile phase101remained at 100% acetonitrile for a further 50 mm before being changed back to 55%acetonitrile in water for the next analysis. The concentration of cis-4,5-dihydroxy-4,5-dihydropyrene in each sample was obtained from the peak area measurement of the peakat a retention time of 10.9 mm calibrated with that of standard cis-4,5-dihydroxy-4,5-dihydropyrene. These procedures were also used to analyze the extracts of the pore watersamples that were obtained by forcing the interstitial water from the wet sediment under20 psi nitrogen and filtering through a 0.4-pm filter. Approximately 1 L of each porewater sample was extracted by using three subsequent washes of ethylacetate (200 ml),and the extract was concentrated to approximately 0.5 ml prior to the HPLC analysis.4.3. Results and Discussion4.3.1. Mineralization of pyrene by indigenous microorganisms in sedimentsMineralization of pyrene to CO2 and H20 was assessed by measuring the amountof 14C02 produced from a culture containing a known amount of14C-4,5,9,10-pyrene.Since 14C-pyrene was not uniformally labelled, the amount of 14C02 detected is anindirect measurement of complete mineralization. A known amount of 14C-4,5,9,10-pyrene (20 nCi) was added into a slurry containing sediment and water from the samesite, and this mixture was incubated under ambient temperature for 23 days after whichtime the 14C02 content was determined. From the 14C02 measurements, the percentageof 14C labeled pyrene mineralized was obtained; the results are summarized in Table 4.2.No 14C02 was detected in any of the three controls containing the 14C labeled pyreneand an autoclaved slurry of sediment and sea water, confirming that mineralization ofpyrene does not take place without microbial activity. However, 14C02 was detected inall the six culture samples and was the result of the mineralization of14C-labeled pyreneby the microorganisms present in these samples. It is important to note that noenrichment process was carried out and no additional nutrients were provided to the102culture. The microorganisms and the nutrients involved in these studies exist in thenatural samples obtained from Kitimat Ann. The present results clearly demonstrate themineralization of pyrene under conditions similar to those in the natural environment,suggesting that the mineralization process likely takes place in the Kitimat Armenvironment.Table 4.2. Pyrene concentration and 14C02 produced from the mineralization of14C-4,5,9, lO-pyrene in indigenous sedimentsSample Concentration of pyrene 14C02,% producedjig/g (dry weight)Controls (autoclaved)CD-i 0CD-3 0CD-6 0SedimentsCD-i 34.4 11±4CD-2 18.1 3±2CD-3 0.6+3.3* 11±2CD-4- 6±2CD-5 1.2 2±2CD-6 3.0 28±2*original pyrene in sediment was 0.6 pg/ml and 3.3 jig/g was added in the laboratory-not evaluatedThe results in Table 4.2 also show that the amounts of 14C02 produced rangefrom 2% to 28% depending on the samples used in the cultures. These samples wereobtained from six locations as shown in Figure 4.1. This variation of minerlizationefficiency cannot be ascribed to a single cause. A number of factors including the103concentration and availability of pyrene; the population of pyrene-degradingmicroorganisms; and the characteristics of sediment samples such as organic contentsmay all contribute to the difference in mineralization.The concentration of PAHs in the surface sediment in Kitimat Arm has beenreported in several studies (64, 65, 66, 72) and the spatial distribution has been discussed(72). It was reported (64, 72) that sediment samples near shore, including the locationsnear our sampling sites, contain elevated concentration of PAHs because of theirproximaty to the source, the Alcan aluminum smelter. The results obtained in ourlaboratory (65, 66) on the PAH concentrations in the near shore sediment samples, CD1-CD6, are summarized in Table 4.1 and Table 4.2. The total concentration of PAHs is inthe range of 6.7 to 527.8 p.glg (dry weight), and the concentration of pyrene is 0.6 to 34.4p.g/g. Thus the microorganisms present in this environment are exposed to elevatedconcentrations of pyrene, which may help them to develop an ability to degrade pyrene.The results in Table 4.2 indicate that there is no direct correlation between pyrenemineralization efficiency and the concentration of pyrene. It is probably moreappropriate to consider the available pyrene concentration instead of the total extractablepyrene concentration in the sample. However this information is not available.The presence of other components in the sample may also affect the microbialmineralization of pyrene. Some microorganisms tend to use easily-degradable, smallerorganic compounds as their carbon and energy sources for growth (94, 192). As aconsequence, pyrene would not be degraded because it is more difficult to break down.The reason for the low mineralization rate in sediment CD2 is not known. This may bepartially due to its higher organic matter content, reducing the availability of pyrene tomicroorganisms. The organic content in sediment CD2 was 3.1%, and all the othersediment samples obtained throughout Kitimat Arm generally have organic contentbelow 2 % cexcept for the sediments obtained from the Yacht Club that haveapproximately 4% (66).104Sediment CD6, a sandy material, was found to give the highest mineralization ofpyrene. This is probably because of (i) relatively high concentration of available pyreneand (ii) a lower content of other organics.Enhancement of the mineralization of pyrene in the sediment samples was studiedby introducing additional microorganisms into the sediment cultures. A mixed culture,Pysed-1, was used. This is a pyrene-enriched culture which originated from sedimentsample C-sed obtained from the location very close to CD 1. This culture has the abilityto mineralize pyrene as discussed in Section 3.3.2. The Pysed-1 culture (1 ml), and 14Clabeled pyrene was incubated in a slurry of sediment and seawater for 23 days under theambient temperature (8-15 °C) in our laboratory and the 14C02was measured. Samplesfrom three locations, CD1, CD3, and CD6 were studied. The percentage of pyrenemineralized was obtained from the 14C02 measurement and data are summarized inTable 4.3. No mineralization appeared in the autoclaved sterile controls, in agreementwith earlier results in Table 4.2. However, in cultures containing both the indigenousmicrobes and the introduced microbes, significant amounts of pyrene were mineralized,approximately 22%, 17%, and 29% from cultures containing sediment CD1, CD3, andCD6, respectively. The addition of the Pysed-1 culture resulted in a 100% enhancementin the mineralization of pyrene in the sample CD1 relative to cultures with onlyindigenous microbes descrdibed in Table 4.2 and suggests a potential use of the enrichedculture in bioremediation. Approximately 54% enhancement of mineralization wasobtained in CD3 by the addition of Pysed-1, and very little enhancement (3.5%) wasobserved in CD6. The reason for this discrepancy is not understood; however, differencesbetween the introduced culture Pysed-1 and indigenous microbes in sediment CD1, CD3,and CD6 may account for some of this difference.105Table 4.3. Enhancement of the mineralization of pyrene by the addition of culture Pysed-1Sample name Autoclaved control Sediment with Pysed- 1 Enhancement, %14co2,%CD-i 0 22±6 100CD-3 0 17±2 54CD-6 0 29±2 Determination of cis-4,5-dihydroxy-4,5-dihydropyrene, a metabolite from pyrenedegradation, in the natural environmentThe results described above, concerning the mineralization of pyrene bymicroorganisms present in untreated sediment, suggest that microbial degradationprocesses likely take place in the environment. Further studies were carried out tosupport this contention because the presence of a degradation metabolite would be strongevidence for degradation. Therefore, a considerable effort was made to identify cis-4,5-dihydroxy-4,5-dihydropyrene within the natural samples.The pyrene degradation metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene, waspreviously studied, as illustrated in Figure 3.8. The metabolite is further mineralized togive CO2 and H20. However, only after the pyrene is degraded to very low level does106the metabolite concentration gradually decrease to undetectable levels (Figure 3.6).Therefore, there is the possibility that cis-4,5-dihydroxy-4,5-dihydropyrene is present at adetectable level if there is a continuous supply of pyrene available for microbialdegradation. It is known that PAHs are constantly released from the aluminum smeltingplant to the Kitimat environment and there is no known natural source of cis-4,5-dihydrodiol pyrene. Thus the presence of this compound would be strong evidence of themicrobial degradation of pyrene in the field.Figure 4.2 (a) shows a chromatogram of an acetonitrile extract obtained from asediment sample, STN2. A fluorescence detector was used for detection with excitationand emission wavelengths of 260 nm and 370 nm, respectively. A similar chromatogramwas also obtained from this sample when UV detection at 254 nm was used. The smallpeak at retention time 10.9 mm (Figure 4.2a) corresponds to that of the standard, cis-4,5-dihydrodiol pyrene (Figure 4.2b). Analysis of the co-injected sample and standard, at 1:1ratio, shows a single peak eluted at 10.9 mm (Figure 4.2c). From the peak height andpeak area measurement of the peak at 10.9 mm, the recovery of the spiked standard isfound to be approximately 95%. The chromatograms in Figure 4.2 demonstrate that theHPLC retention time of the compound of interest in the sample matches that of cis-4,5-dihydrodiol pyrene.The fraction of 10.9 mm retention time was collected after eluting from both thepreparative and analytical HPLC columns, and was further characterized by usinganother HPLC column (ODS-2, from GL Scientific). This general purpose reversedphase C18 column more strongly retains the compound, giving a retention time of 25.6mm. Standard cis-4,5-dihydrodiol pyrene also elutes at 25.6 mm in agreement with theresult of Heitkamp and Cerniglia (101). A mixture of the sample and the standard (1:1ratio) also eluted at the same retention time, suggesting that the compound in the samplematches the standard.107‘ J\\,)\(b)/\ (c)t \ 1) L \ / \ / II I I0 4 8 12 16 20 24 28Retention Time, mmFigure 4.2. Chromatograms obtained from (a) sediment sample STN2;(b) cis4,5dihydroxy-4,5-dihydrOpyrefle standard; and (c) 1:1 mixture of (a)and (b).108As mentioned in the previous Chapter, a direct UV/visible scan of the sample didnot provide useful information because of the insufficient sensitivity. In addition, theimpurities in the fraction made it difficult or even impossible to obtain a UV/visiblespectrum corresponding to the trace amount of the compound of interest. Therefore aseries replicate HPLC/UV determinations of the same sample were carried out, onlychanging the wavelength of the UV detector after each determination. By plotting thepeak intensity vs. wavelength, a UV/visible spectrum can be obtained. Figure 4.3 showsspectra obtained from the sample and cis-4,5-dthydrodiol pyrene. Similar absorptionmaxima were observed. The identical HPLC retention time and similar absorptionspectra between the compound and cis-4,5-dihydrodiol pyrene strongly suggest that thecompound of interest is cis-4,5-dihydrodiol pyrene, one of the major metabolites fromthe microbial degradation of pyrene.GC/MS was used to further confirm the indentification of the separatedcompound. Sediment sample STN2 underwent extraction and preparative columnseparation. The fraction collected from the preparative column was acetylated, asdescribed in Section 4.2.3. The chromatogram and mass spectrum of the acetylationproducts analyzed by GC/MS are shown in Figure 4.4. It is clearly observed that thecompound isolated from the sediment STN2 has identical retention time (Figure 4.4 a) tothat of the standard shown in Figure 3.11(a). The mass spectrum of this compound iscomposed of M peak at mlz 320, and fragment ions at m/z 260, 218, and 189, whichresembles that of the standard cis-4,5-dihydroxy-4,5-dihydropyrene (Figure 3.11(b)).These results strongly support the presence of cis-4,5-dihydroxy-4,5-dihydropyrene in thesediment sample STN2. Therefore, the degradation of pyrene in the natural environmentappears very likely.1094Ct=—1• I • • •200 220 240 260 280 300 320Absorption Wavelength, nmFigure 4.3. Comparison of UV absorption spectra obtained from(• ) cis.dihydroxy-4,5thYdr0PY standard; and(0 ) purified fraction from sediment sample STN2.%age0. 28:00 30:00‘- ‘‘1’’’32:00 34:00 36:00218.0RT(b)49.0083.90 50 00260.0319.9150 300 380 400 mfzFigure 4.4. GC/MS trace (a) and mass spectrum (b) of the acetylation product of themetabolite isolated from the sediment sample STN2(Note: the mass spectrum of the acetylated cis4,5dihydroxy4,5-dihYdr0PYrefle standardshown in Figure 3.llb)111A number of sediment samples from various locations were analyzed by usingHPLC/fluorescence detection. The results are summarized in Table 4.4. Sedimentsamples obtained from near the shore of the Kitimat Yatch Club including STN2, C-Sed,and CD1 were found to contain approximately 10-20 nglg (dry weight) of cis-4,5-dthydroxy-4,5-dihydropyrene. Other sediment samples obtained further away from theshore in Kitimat Arm contained much lower concentrations, some undetectable, of thiscompound. The detection limit for this determination was 0.5 ng/g for a 10 g sedimentsample and 5 pg/ml for 1 L water sample. A detectable amount (0.5 ng/g) of dihydrodiolpyrene was measured in K5a-A. No dihydrodiol pyrene was detected in the sedimentsample from K5a-B and K5a-C. Generally, relatively higher concentrations ofdihydrodiol pyrene are found in the sediments that contain higher concentrations ofpyrene.No dihydrodiol pyrene was detected in sea water samples(5 pgIL). This may bebecause pyrene tends to accumulte into sediment because of its hydrophobicity. A smallamount of cis-4,5-dihydroxy-4,5-dihydropyrene was found in pore water samples whichwere obtained from wet sediments CD1 and CD3 (Table 4.4). The concentration ofdihydrodiol pyrene was lower than that in the sediment from the same site, probablybecause the solubility of cis-4,5-dihydroxy-4,5-dihydropyrene is lower in water than inorganic solvents, and tends to associate with sediment. We have observed that thiscompound is more soluble in methanol than water.The present study provides the first evidence of the presence of cis-4,5-dihydroxy-4,5-dihydropyrene in a natural environment and supports the presence ofmicrobial degradation of PAHs in Kitimat Arm. Laboratory cultures clearly demonstratethe degradation and potential mineralization of pyrene. In order to relate laboratorystudies to the natural environment, a number of environmental factors need to beconsidered. The effect of some of these factors on the degradation of pyrene aredescribed in the following sections.112Table 4.4. Concentration of dihydrodiol pyrene in samplesSample name Concentration of dihydrodiol pyrene, ng/gSedimentBlank ndSTN2 20 (dry weight)Csed* 10—20 (dry)CDi*l 1 (wet)CD3*l—O.5(wet)K5a-A detectable (dry)K5a-B ndK5a-C ndKA-7 ndPore water ng/mlCD-i -1CD-3 <CD-iWater columnSTN4 (surface) ndk5A (surface) ndK5A (at 50 m) nd*: the sediment used as inoculants for generating cultures used in Chapter 2 and 3* 1: the sediment after pore water was squeezed out113Detection limit: 0.5 nglg4.3.3. Effect of temperatureTemperature is an important factor which can influence the growth ofmicroorganisms and therefore affect the microbial degradation of pyrene. Thedegradation studies discussed above were carried out at a constant temperature of 26 °C.However, the temperature in the natural environment varies and can reach 4 °C on theocean floor. In order to obtain information on the effect of temperature, experiments onthe microbial degradation of pyrene were also carried out at lowered temperatures,namely 10 and 2 °C. Figure 4.5 shows pyrene degradation results obtained when cultureswere maintained at 26 °C (curve a), 10 °C (curve b), and 2 °C (curve c), compared withthe control which was conducted at 26 °C (curve d). Clearly no significant loss wasobserved in the control. However the degradation of pyrene by microorganisms wasobserved at all three temperatures. The rate decreases as the temperature decreases. Whenthe degradation was carried out at 26 °C, 10 .ig of pyrene in the culture was degraded toan undetectable level (<10 ng!ml) within 200 hr. After the same period of incubationtime, 40% and 70% of the pyrene were left in the cultures that were maintained at 10 °Cand 2 °C, respectively. Although the rate of degradation of pyrene is reduced at lowertemperature, degradation does take place at 2 °C. Thus the microorganisms obtained inthis study are able to grow at these low temperatures and maintain their ability todecompose pyrene. These results indicate that bioremediation could be possible underlow temperature conditions.11410- 8>.l- 600c-)20Incubation time, hrFigure 4.5. Effect of incubation temperature on the microbial degradation of pyrene.(0 ) uninoculated control at 26 °C; (• ) culture at 2 °C; (S ) culture at 10 °C; and(A ) culture at 26 0C0 200 400 600 800 10001154.3.4. Effect of culture medium and tolerance of saltThe make up of the culture medium has a significant effect on the phenanthrenedegradation as discussed in Chapter 2, and therefore the effect of medium on thedegradation of pyrene was also investigated in the present study. Strain Pysed-1-Y wasused as inoculant to degrade pyrene, and experiments with the presence and absence ofFe3+ in the culture medium were compared. While Fe3+ has a profound effect on thedegradation of phenanthrene as discussed in Section 2.3.4, little difference was observedon the degradation of pyrene. These results suggest that the enzyme systems involved inthe degradation of phenanthrene and pyrene may be different.On the other hand, the addition of yeast extracts to the culture medium resulted inan enhanced degradation of pyrene (Figure 4.6), similar to that observed forphenanthrene. Yeast extract is rich in organic nutrients, which enhanced the growth ofmicroorganisms as shown in Figure 3.1. In addition to the enhancement in growth anddegradation, yeast extract also increased salt tolerance of the microorganisms. In thepresent study, salt tolerance of the microorganisms was examined by the addition ofNaCl (3%) into the mineral salt medium.Data on the degradation of pyrene in the presence of various additionalcomponents in the culture medium are summarized in Table 4.5. In the mineral saltmedium, fast degradation was observed, thus 10 ig of pyrene was completely degradedto undetectable level after 90 hr of incubation. The degradation of pyrene was slowerwhen 3% NaCl was present in the mineral salt medium. However, when other organicnutrients, such as yeast extract and peptone, were added, the inhibition of pyrenedegradation was significantly reduced as shown in Table 4.5. The complete degradationof pyrene was achieved when yeast extract or peptone was present in the culture medium,while 72% of pyrene still remained in the culture when pyrene was incubated in themineral salt medium containing 3% NaC1 for 137 hr. These results are consistent withthose reported earlier on the degradation of phenanthrene.116C,)zC.)C0L)Incubation time, hrFigure 4.6. Effect of culture medium on the degradation of pyrene by culture Pysed-1-Y(C) ) mineral salt medium control(• ) culture in mineral salt medium(A ) culture in mineral salt medium without Fe3( ) culture in mineral salt medium and yeast extract (250 jag/mi)(D ) mineral salt medium and yeast extract control.121086400 40 80 120117Table 4.5. Percentage of pyrene left in cultures when different medium compositions areused250 ppm peptoneand 3% NaC1 in M.SInoculant: Phensed-2, 0.5 mlInitial content of pyrene (at time 0): 10 pgnd: not detectable ( <2 nglml or 20 ng in culture)4.3.5. Effect of pyrene concentration and microorganism populationAs discussed in Chapter 2, the rate of phenanthrene degradation by mixedmicroorganisms is dependent on the concentration of phenanthrene and the population ofthe microorganisms. Therefore, detailed studies on the possible effect of concentration ofpyrene and population of microorganisms were also carried out.Three sets of experiments involving the incubation of different amounts of cultureinoculant (0.1 ml or 0.5 ml) and pyrene (10 jig or 100 jig) were conducted, and theresults on pyrene degradation are shown in Figure 4.7. Unlike for phenanthrenedegradation, where the degradation rate increases with the amount of culture inoculantMedium Incubation time, hr.0 90 137 162Uninoculated control 100±2 91±10 90±4 ±90Mineral salt (M.S.) 100±2 nd nd nd3% NaC1 in M.S. 100±2 88±0 72±5 27250 ppm glucoseand 3% NaClinM.S. 100±2 71±25 42±1 1250 ppm yeast extractand 3% NaCl in M.S. 100+2 56±5 nd nd100±2 21±1 nd nd118used (Figure 2.3), the present study shows that increasing the culture inoculant from 0.1ml to 0.5 ml has little effect on the pyrene degradation. This is probably becausemicroorganisms tend to utilize the dissolved PAils (237, 238), and the degradation ratein the present case is limited by the concentration of pyrene dissolved in the culturemedium. The solubiity of pyrene in aqueous medium is only 0.14 jig/mi as compared to1.3 jig/ml for phenanthrene (106). Thus the solubility is probably a more importantlimiting factor for the degradation of pyrene than for phenanthrene.An increase of the amount of pyrene in 10 ml water from 10 jig to 100 jig wouldnot increase the concentration of pyrene in the solution because it would reach saturationin both cases. However, in the present dynamic situation where the dissolved pyrene iscontinuously being utilized by microorganisms, an increase of pyrene in the cultureresults in an increase of the total pyrene degraded (Figure 4.7). The limiting factor isprobably also the low solubility of pyrene.These results suggest that the degradation of indigenous pyrene associated withsediment would be limited by the desorption process. It is important to carry outmicrobial degradation (mineralization) studies by using indigenous sediment as discussedin Section 4.3.1, in order to know the biodegradability of pyrene in a specific sedimentsample. Without this information, it is impossible to assess the persistence and the fate ofpyrene (or other pollutants) in the environment.The degradation of pyrene was also studied by measuring the metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene. Figure 4.8 show the amount of the metabolite present atvarious incubation times. Similar patterns of metabolite production were obtained when10 jig pyrene was incubated with 0.1 ml or 0.5 ml culture inoculant. When the amount ofpyrene is increased from 10 jig to 100 jig, approximately 3 fold increase of metabolite isobserved. In all three cases, the maximum amount of the metabolite was about 5 to 10%of the pyrene degraded. The maximum appeared in the cultures when pyrene wasdegraded to less than 50% of its initial concentration. The metabolite, as a result of the119Cr-)initial oxidation of pyrene, remained in the culture only for a short period of time, andwas further degraded.1210864201201008060402005000 100 200 300 400Incubation time, hrFigure 4.7. Effect of initial pyrene concentration and the amount of culture on thedegradation of pyrene.(•) 10 jig pyrene, 0.5 ml culture() 10 jig pyrene, 0.1 ml culture(A) 100 jig pyrene, 0.5 ml culture (right Y-axis)(0)10 jig pyrene, control.• 1205-4.3.2C.)— 1CC.)C ACL) • •0 100 200 300 400 500Incubation time, hrFigure 4.8. Amount of cis-4,5-dihydroxy-4,5-dihydropyrene in culture at various intervalsof incubation.Legend as denoted in Figure 4.7.1214.3.6. Degradation of pyrene in the presence of cis-4,5-dihydroxy-4,5-dihydropyrene orpyrenolPyrene in the real environment is accompanied by many other compounds andtheir presence can have profound impact on the fate of pyrene with respect to microbialdegradation. Thus degradation of pyrene in the presence of other biodegradablecomponents was studied. Typical metabolites from pyrene degradation, cis-4,5-dihydroxy-4,5-dihydropyrene and pyrenol, were used as examples to assess their effect onthe degradation of pyrene.Figure 4.9 shows the amount of pyrene and cis-4,5-dihydrodiol pyrenedetermined in the cultures at various incubation times. The degradation of pyrene in theabsence and presence of additional 7.8 tg of cis-4,5-dihydroxy-4,5-dihydropyrene wascompared. As shown in Figure 4.8, the degradation of pyrene is relatively slower whenthe additional cis-4,5-dihydroxy-4,5-dihydropyrene is present in the culture. The amountof cis-4,5-dihydroxy-4,5-dihydropyrene in the culture at any given time is the net result ofthe combination of 7.8 ig cis-4,5-dihydroxy-4,5-dihydropyrene that was originally addedinto the culture, the production of the metabolite from the degradation of pyrene, and thefurther degradation of the metabolite. It is clear that the degradation of pyrene preceedsthat of the metabolite, cis-4,5-dihydroxy-4,5-dihydropyrene. A significant decrease of cis4,5-dihydroxy-4,5-dihydropyrene is obtained only after pyrene in the culture is reduced toless than 40% of its amount in the initial culture. These results are consistent with thosediscussed earlier when no metabolite was added into the culture. These results all indicatethat the microorganisms obtained in this study utilize pyrene faster than cis-4,5-dihydroxy-4,5-dihydropyrene. This explains why cis-4,5-dihydroxy-4,5-dihydropyrene isobserved in all pyrene degradation cultures at the appropriate incubation time and innatural samples from Kitimat Arm, where the microbial degradation of pyrene isexpected to take place.12212zL)__________________0 100 200 300 400Incubation time, hrFigure 4.9. The amount of pyrene and cis-4,5-dihydroxy-4,5-dihydropyrene in cultures,measured at various intervals during incubation.(0 ) pyrene in control(A ) pyrene in the culture, without the addition of cis-4,5-dihydroxy-4,5-dihydropyrene(• ) pyrene in the culture, with the addition of cis-4,5-dihydroxy-4,5-dihydropyrene() cis-4,5-dihydroxy-4,5-dihydropyrene in culture(D ) cis-4,5-dihydroxy-4,5-dihydropyrene in control.123In another experiment, the degradation of pyrene in the absence and presence of 3jig and 30 jig of pyrenol was carried out. The amount of pyrene in the cultures measuredat several intervals during incubation is shown in Figure 4.10. When no pyrenol wasadded into the culture, pyrene was rapidly degraded to less than 5% after 200 hr ofincubation. However, no significant degradation of pyrene was observed during the sameperiod of incubation when 3 jig of pyrenol was added into the culture. Further incubationof this culture to 400 hr results in the degradation of pyrene to approximately 10% of itsinitial amount. The additional 30 jig of pyrenol into the culture dramatically reduces thedegradation of pyrene. Little pyrene was degraded when the culture containing 10 jigpyrene, 30 jig pyrenol, and 10 ml mineral salt medium, and 0.1 ml of an enriched culturePysed- 1 was incubated for as long as 450 hr. This clearly demonstrates that pyrenolinhibits the degradation of pyrene. This is probably because the microorganismspreferentially uptake pyrenol when both pyrene and pyrenol are present. This point issupported by the results shown in Figure 4.11. The contents of pyrene and pyrenol incultures at various intervals during incubation are compared, as shown in Figure 4.11.While little reduction of pyrene concentration is observed, pyrenol (30 jig) is reduced toless than 50% after 450 hr of incubation. The preferential removal of pyrenol to pyreneby the microorganisms probably explains why pyrenol was not observed during pyrenedegradation. These results on the degradation of pyrene and its metabolites supportearlier finding reported in this thesis on the presence of cis-4,5-dihydroxy-4,5-dihydropyrene in cultures and in samples.12412zzC) 8.-C-.‘C0500Incubation time, hrFigure 4.10. The amount of pyrene in cultures and in the control, measured at variousintervals during incubation.(• ) without the addition of pyrenol(A ) with the addition of 3 ig of pyrenol( ) with the addition of 30 jig of pyrenol(0 ) sterile control0 100 200 300 400125L.-..100::I 500Incubation Time, hrFigure 4.11. Relative concentration of pyrene ( •) and pyrenol ( B ) in cultures at variousincubation times compared to those in the culture at incubation time 0.0 100 200 300 400126Chapter 5. Degradation and Determination ofBenzo(a)pyrene and the Mixed PAHs5.1. IntroductionBenzo(a)pyrene (Bap) is a well known carcinogen, and it is found to be present inthe Kitimat Ann environment at a concentration as high as 56 mglg in sediment samples,as discussed in Section 4.2.1. Although the metabolism of Bap in mammals, yeast, andfungi, which is a monooxygenase pathway, has been well studied (94, 105), very little isknown about the bacterial degradation of this PAH. This is because PAHs with four ormore aromatic rings are generally found to be resistant to bacterial degradation. However,because the microorganisms isolated in this thesis work, as described in the previouschapters, show an ability to degrade both phenanthrene and pyrene, these microorganismswere used in studies of the degradation of higher molecular weight PAHs including Bap.Microbial degradation studies are often carried out by using a single PAH as thetarget compound in the cultures, because a well defmed system provides usefulinformation about PAH degradation including metabolism pathways. However, in realityPAHs are usually found in environmental samples as a complex mixture of a large numberof components: many of the sixteen PAHs listed by the US EPA as priority pollutants areoften present (1, 2, 6, 164). Little is known about the microbial degradation of mixtures ofPAHs. Therefore in the present work some preliminary studies had been carried out toexplore the degradation of the mixed sixteen PAHs by using the microorganisms obtainedfrom the Kitimat samples. Such a study should be useful for an assessment of the fate ofPAHs on the Kitimat environment.1275.2. Experiment5.2.1. ChemicalsBenz[a]pyrene (99.99%) was obtained from Aldrich. A standard containing thesixteen PAHs as shown in Figure 1.1 was purchased from Supelco: the solvent is amixture of benzene and methanol (1:1). The working solutions for degradation studieswere prepared by diluting the commercial standard with acetone. For GCIFTD analysis, thestandard was prepared in toluene. All solvents used were HPLC grade and were purchasedfrom Fisher Scientific.The culture media used for the degradation studies include the mineral salt medium(M.S.), and M.S. containing Tween-80 (200 mglml) or peptone and yeast extracts (250mglml). These media were prepared as described in Section GC/FID analysisA Hewlett Packard gas chromatograph Model 5890 equipped with a flameionization detector (FID) was used for the determination of PAHs. A capillary column (30m x 0.32 mm i.d., 0.25 mm coating, PTE5 from Supelco) was used to separate thecomponents in the samples. A sample (1 ml) was injected into the GC injector (260 °C,splitless mode). Helium (70 mI/min) was used as carrier gas and nitrogen (30 mI/min) asmake-up gas. The GC oven temperature was programmed as follows: holding at an initialtemperature of 100 °C for 4 min, then ramping at 5 °C/min to 200 °C and holding for 3min, followed by ramping at 8 °C/min to 250 °C and holding for 5 mm, and fmallyramping at 12 °C/min to 300 °C and holding for 30 mm to elute all the components out ofthe column before returning to the initial temperature for the next analysis. The FID wasoperated at an air flow 300 mI/mm and a hydrogen flow 30 mI/mm. The detector wasmaintained at 320 °C. GC traces were recorded by using a HP 3393A integrator and peakareas of the chromatographic signals were used for quantification by calibration againstthe internal standard.1285.2.3. Degradation and determination of Bap and the sixteen PAHsThe mixed culture Pysed-1 was used as inoculant for the degradation studies ofBap and the sixteen PAHs. The experiments were performed as described in Section 3.2.7except that Bap or the sixteen PAHs were used instead of pyrene. At the end of thedesired incubation time 20 ml of acetonitrile was added into the flask and the supernatantof the culture was analyzed for Bap by using HPLC/fluorescence. The HPLC analysis wascarried out by using a Supelcosil1LC-PAH column and 100% acetonitrile as mobilephase flowing at 1 mI/mm. The fluorescence excitation wavelength was set at 384 nm andemission wavelength 404 nm.For the determination of the sixteen PAHs, the cultures were extracted by usingthe following procedures. A known amount (20 ml of 1450 mg/mi) of hexamethylbenzenewas added to the culture as an internal standard and then the contents of the culture wereextracted with 5 ml of CH2C12, three times. These three fractions were pooled togetherand the solvent was evaporated to dryness under a gentle N2 stream. The residue wasdissolved in 2 ml of toluene and the solution was subjected to GC/FID analysis.5.2.4. Extraction and clean-up of the sediment samplesEight portions of the sediment sample (C-sed) that was used to generate thecultures for this thesis work, were freeze-dried and the water content in the sediment wasdetermined to be 66% (wlw). The eight dry sediment samples were separately extracted asfollows. The dry sediment was ground to fme powder prior to the extraction. A portion ofthe powder (10 g) was accurately weighted and placed in a Soxhlet thimble together with20 ml of the internal standard, hexamethylbenzene (1450 mg/mi). Approximately 2 g ofdry Na2SO4 was added on top of the sediment powder to prevent the sample fromfloating out of the thimble. Soxhiet extraction was then carried out by using 150 ml ofCH2C12 in a 250-ml round-bottom flask. The flasks were heated to maintain the solvent at129reflux. The top of the water cooled condenser was connected to a drying tube containingdry Na2SO4. The extraction was stopped after 7 hours. The extract was condensed toabout 1 ml by using a rotary evaporator over a water bath at 30 °C. Approximately 2 g ofdry Na2SO4 was added to each condensed extract and the extract residue was transferredonto the Na2SO4 by evaporating the solvent.A clean-up column was prepared by packing 3 g of Florisil (2% deactivation) intoa glass column (30 cm x 12 mm i.d.) fitted with a glass wool plug on the bottom. The dryNa2SO4 that had adsorbed the sample extract was added to the top of the Florisil column.Some Na2SO4 particles trapped on the column wall were washed down by using 1 ml ofhexane. The elution was carried out by using 15 ml of hexane followed by 25 ml of amixture of hexane and dichioromethane (1:1). The first 8 ml fraction of hexane eluent thatis mainly saturated hydrocarbons was discarded. The rest of eluent was collected forfurther GCIFID analysis.The eluent was condensed to about 1 ml by using the rotary evaporator. Thiscondensed fraction was transferred to a small sample vial and 0.5 ml of toluene was added.This solution was further evaporated to remove dichloromethane and hexane. The fmalsolution was prepared in toluene (approximately 1 ml) and used for GC/FID analysis asdescribed in Section 5.2.2.In order to determine the loss of PAHs over time, duplicate samples of sedimentCD-i were stored in Erlenmeyer flasks covered with cotton plugs, and the flasks wereplaced on a shaker (136 RPM) for a month at ambient temperature. Duplicate controlsampies were prepared by autoclaving two other portions of sediment CD-i twice at 120°C: they were also kept on the shaker for a month under the same conditions. The sampleswere subsequently freeze-dried, extracted, cleaned-up, and analyzed by using the sameprocedures as described above.1305.3. Results and Discussion5.3.1. Degradation of benzo(a)pyrene (Bap) by the enriched culturesA preliminary experiment to assess the possible degradation of Bap as sole carbonsource was carried out in a culture containing Pysed-1 in the mineral salt medium. Theconcentration of Bap in the cultures and in the sterile controls was monitored by using theHPLC/fluorescence detection system. The concentration of Bap, relative to its initialconcentration, at the incubation times 30 and 64 days is shown in Table 5.1 (Row I). Only46% of Bap was detected (54% degraded) in the cultures after 64 days of incubation. Noloss of Bap was observed in the uninoculated controls, thus the depletion of Bap in thecultures containing Pysed- 1 can be attributed to microbial degradation. There are very fewexamples of the microbial degradation of PAHs as large as Bap (106, 124, 205, 239) andthe reported microbial degradation of Bap (124, 205, 239) involved other carbon sources.It is clear that the cultures generated from the Kitimat samples have remarkable properties.Similar results were also obtained when the culture Pysed-2 was used as theinoculant. The cultures Pysed-1 and Pysed-2 originated from a sediment sample, andpyrene was used as the sole carbon source during the enrichment process. The presentresults and those in Chapter 3 show that these enriched cultures are capable of degradingboth pyrene and Bap separately. Further experiments were carried out to investigate thedegradation of these two PAHs when they were both present in a culture and to study theeffect of pyrene on the degradation of Bap. Both pyrene and Bap, each at 1 mg/mi level,were added into a culture of Pysed-1. The relative concentration of the Bap after 64 daysof incubation is 33% as shown in Row II of Table 5.1, but pyrene is not detected. Thuswhen both pyrene and Bap are present in the culture, pyrene is completely degraded toundetectable level prior to Bap. These results show, as expected, that Bap is more difficultto degrade than pyrene for this mixed culture. However, it is possible that because the131mixed culture Pysed-1 was enriched with pyrene, more pyrene-degraders were present inthe cultures, leading to faster degradation of pyrene.Table 5.1. Percentage of benzo(a)pyrene remained in the culture (Pysed-1) after 30 and 64days of incubation relative to the initial concentration of benzo(a)pyreneExperiment Medium Incubation Time (day)No. Composition 30 64 64Bap, % Pyrene, %I MS+Bap control* 100±1 102±2culture 91±8 46±8II MS+Bap+Py control- 99±2 100±2culture- 33± 10 0III MS+Bap control 104±3 93±9+Peptone+Yeast extract culture 72±2 43÷7IV MS+Bap control 47±5 39± 10+Tween-80 culture 20±14 13±6MS: mineral salt medium. Bap: benzo(a)pyrene Py: pyrenePeptone and yeast extract each at 250 mg/ml concentrationTween-80 at 200 mg/mi concentration-: not analyzed because of loss of samples*: uninoculated control**: standard deviationFurther studies were carried out in attempts to improve the rate of microbialdegradation of Bap. The results presented in Chapter 4 demonstrated an enhancement ofpyrene degradation following the addition of organic nutrients, such as peptone and yeast132extracts, to the cultures. Because of this success, organic nutrients were also added to Bapcultures. The relative concentrations of Bap in the organic nutrient supplemented culturesand the controls are summarized in Table 5.1 (Row III). A comparison of the results inRow III with those in Row I reveals that the degradation of Bap is enhanced initially bythe addition of Peptone and yeast extracts. The relative concentration of Bap at theincubation time 30 days is reduced to 72% in the presence of organic nutrients, whereaslittle degradation is observed in the same incubation period without organic nutrients inthe cultures. When the incubation time is increased to 64 days, there is little difference inthe extent of Bap degradation in the cultures with or without peptone and yeast extracts.It is likely that the organic nutrients help the microorganisms to adapt to the new mediumand to grow faster initially, as is indicated by the higher turbidity of the cultures. As aresult, the degradation of Bap is enhanced at the earlier period of incubation time. As theincubation time is increased, both cultures have sufficient time to reach a growthmaximum and to degrade similar amount of Bap. No loss was observed in the sterilecontrols again indicating that the degradation in the cultures is due to microbial activity.As discussed in Chapter 4, the dissolution of pyrene is possibly the rate limitingstep for pyrene degradation. Bap has a solubility of 3 mg/L (106) which is over 40 timeslower than the solubility of pyrene (129 mgIL). Clearly, the limited solubility of Bap inwater could affect its degradation. In order to investigate this, a surfactant, Tween-80(200 mglml), was added to the cultures and the controls in order to improve the solubilityof Bap. Surprisingly, the relative concentration of Bap in both the controls and thecultures, measured by using the HPLC/fluorescence system, is decreased, Row IV ofTable 5.1. However, the loss of Bap in the cultures is greater than in the controls.For comparison, the ratio of Bap in the cultures relative to the controls at a givenincubation time is shown in Table 5.2; the lower the ratio, the more the degradation. Theratios are lower in the presence of Tween-80, (0.42 and 0.33 at the incubation times 30and 64 days, respectively), indicating that Tween-80 enhances the degradation of Bap.133The cultures containing Tween-80 have higher turbidity than those without Tween-80, andno growth was observed in the uninoculated controls. Thus the enhanced degradation mayresult from both the increased solubility of Bap and the improved growth of themicroorganisms. This is consistent with those obtained from pyrene degradation studies.However, further study is necessary in order to understand how surfactants enhance themicrobial degradation of high molecular weight PAHs.Table 5.2. Ratio of benzo(a)pyrene in the culture to that in the controlMedium Incubation Time (day)Composition 30 64MS+Bap 0.9 1±0.08 0.45±0.08MS+Bap+Tween-80 0.42±0.14 0.33±0.10The apparent decrease of Bap in the controls is not fully understood. It is probablythe result of a change in the fluorescence emission spectrum of Bap when it is mixed withTween-80, leading to a lowering of the intensity of fluorescence of Bap in the micellesolution. It is known that the fluorescence emission spectrum of naphthalene is changedwhen naphthalene is mixed with micelles (240).5.3.2. Degradation of a mixture of sixteen PAHsThe studies described in the previous chapters dealt with the degradation of asingle PAH in each culture. However because many PAHs are present in the samplesobtained from the Kitimat environment (66), it is important to gain some informationabout behavior of multi-component system.134The experiments were carried out in the same way as those involving a single PAHexcept that the mixture of the sixteen PAHs (0.1 ml in acetone) was quantitativelyintroduced into the series of sterile flasks. The acetone was evaporated, and the mineralsalt medium was added followed by 0.5 ml of the enriched culture Pysed-1. The controlswere prepared similarly without the culture Pysed-1. Duplicate cultures and controls wereextracted with CH2C12 at the incubation time 30 and 50 days. The PAHs in the CH2C12extracts were quantitatively determined by using GCIFID. The results from triplicatedeterminations of each of the two samples are summarized in Table 5.3. No quantitativeresults for naphthalene, acenaphthylene, acenaphthene, and fluorene were obtainedbecause of loss during the incubation and extraction processes. The extensive loss of theselow molecular weight PAHs during the incubation in soil has been reported by others(102, 241). The results in Table 5.3 demonstrate that the extent of PAH degradation bythe mixed microorganisms generally decreases as the number of aromatic ring increases.The tricyclic PAHs, including phenanthrene and anthracene, are degraded faster thantetracyclic PAHs such as fluoranthene, pyrene, chrysene, and benz(a)anthracene. Somedegradation is observed for the pentacycic PAHs and essentially none for the hexacycicPAHs.Table 5.3 also shows that the degradation rate of PAH is dependent on thestructure when PAHs containing the same number of aromatic rings are compared. Forexample, phenanthrene, an angular molecule, is degraded faster than anthracene which hasa linear structure. The degradation rates for the tetracyclic PAHs are in the order ofpyrene > benz(a)anthracene > chrysene. Pyrene, which has a cluster structure, has thehighest degradation rate among the three tetracyclic isomers. Similar results wereobserved by Bossert and Bartha (124), who correlated this difference in degradation rateto the solubility of the tetracyclic PAHs.135Table 5.3. Contents (mg) of 16 PAHs in the cultures and controls at the given incubation timeComponent Incubation time (day)Control Culture0 30 58 30 58Content of PAH left (mg)Naphthalene nd nd nd nd ndAcenaphthylene nd nd nd nd ndAcenaphthene nd nd nd nd ndFluorene nd nd nd nd ndPhenanthrene 1.2±0.6 0.5±0.1 0.5±0.4 0.2 ndAnthracene 2.0±0.6 0.9±0.2 0.9±0.4 0.5±0.1 0.3±0.1Fluoranthene 3.4±0.6 1.2±0.2 1.3±0.3 0.8±0.3 0.5±0Pyrene 3.2±0.6 1.0+0.2 1.9±0.5 1.0±0.3 0.4±0.2Benz(a)anthracene 3.9±0.4 4.4±0.7 3.9±0.6 2.6±0.6 1.7±0.6Chrysene 3.5±0.2 4.2±0.8 3.4±0.5 2.5±0.4 1.6±0.5Benzo(b)fluoranthene 3.8±0.3 4.5±0.8 4.3±0.7 3.8±0.6 2.9±0.8Benzo(k)fluoranthene 3.6±0.3 4.7±1.1 4.2±0.7 4.1±0.7 3.5±0.3Benzo(a)pyrene 3.9±1.0 4. 1±0.9 3.7±0.9 3.4±0.7 2.2±0.5Dibenz(a,h)anthracene 4.0±0.4 4.8±1.1 4.6±0.8 4.5±0.6 4.4±0.6Indeno( 1 ,2,3-cd)pyrene 3.8±0.4 4.4±0.8 5.2±0.6 4.2±0.6 4.0±0.5Benzo(g,h,i)perylene 3.7±0.1 4.5±1.1 4.6±0.6 4.2±0.2 4.5±0.8nd- not detected.mean ± standard deviation (n=6)136Table 5.4. LIB values and the relative concentrations of 16 PAHs in the cultures comparedto those in the controlsComponent Number of Incubation time (day) L/Bbenzene ring 30 58 value#Relative concentrationNaphthalene 2 nd ndAcenaphthylene 2* nd nd*Acenaphthene 2 nd ndFluorene 2* nd nd -Phenanthrene 3 0.40 nd 1.46Anthracene 3 0.56 0.33±0. 10 1.57Fluoranthene 3* 0.67 0.38±0. 10 1.22Pyrene 4 1.00 0.21±0. 10 1.27Benz(a)anthracene 4 0.59 0.43±0.15 1.58Chrysene 4 0.59 0.47±0.15 1.72Benzo(b)fluoranthene 4* 0.84 0.67±0.19 1.40Benzo(k)fluoranthene 4* 0.87 0. 83±0.07 1.48Benzo(a)pyrene 5 0.83 0.59±0. 13 1.50Dibenz(a,h)anthracene 5 0.94 0.96±0.10 1.79Indeno(1,2,3-cd)pyrene 5* 0.95 0.77±0. 10 1.40Benzo(g,h,i)perylene 6 0.93 0.98±0.10 1.12nd: not detected.* Another cyclic pentene ring not included.#: From Sander and Wise, Adv. in Chromatogr. 1986, 25, 139.137Studies of PAH retention behavior on reversed phase (C 18) liquid chromatographcolumns show that retention times correlate with the shape of the PAH. The ratio oflength to breadth (LIB) has been proposed as a description of the shape of PAHs (157).The larger the L/B ratio is, the longer is the retention time. In the present study, it is foundthat the rates of biodegradation for tricyclic and tetracyclic PAHs are lower with higherLIB value. As shown in Table 5.4, the L/B values for phenanthrene and anthracene are1.46 and 1.57, respectively, and phenanthrene is degraded faster than anthracene. Asimilar correlation is found with the tetracyclic PAHs. As the L/B ratio increases from1.27 for pyrene to 1.58 for benz(a)anthracene, and to 1.72 for chrysene, the percent ofpyrene, benz(a)anthracene, and chrysene degraded after 58 days of incubation decreasesfrom 79% to 57% and 53%, respectively. Benz(b)fluoranthene (L/B 1.40) is also degradedmore than benz(k)fluoranthene (L/B 1.48). These results show that the shape of PAHsaffects their microbial degradability and in turn their persistence in the environment.The microbial degradation of PAHs is dependent on the nature of microorganismsin addition to the structure of PAHs. The mixed culture Pysed-1 originated from asediment sample containing high concentrations of pyrene, fluoranthene and some otherhigh molecular weight PAHs such as Bap. Consequently, the microbial degradation ofthese PAHs as observed in the present study is possibly the result of the chronic exposureof the microorganisms to PAHs in the sediment.5.3.3. Determination of PAHs in sedimentsThe concentrations of the sixteen PAHs in the sediment (C-sed) that was used togenerate the cultures for most of this thesis work were determined by using GCIFID.Eight replicate sediment samples were Soxhiet extracted by using CH2C12. The extractswere analyzed after clean-up on Florisil columns. The mean concentrations of the sixteenPAHs from the eight replicate samples are presented in Table 5.5. The sediment samplescontain high concentrations of PAHs and the concentrations of the higher molecular138weight PAHs are higher than those of lower molecular weight. The fact that PAHdegrading microorganisms are obtained from this sediment sample gives support to theproposal that the ability of the microorganisms to degrade some of the higher molecularweight PAHs such as pyrene and Bap is associated with chronic exposure to these PAHs.Table 5.5. The mean concentrations of the sixteen PAHs in the sediment sample C-sedMean concentration of PAH (mg g1) RSD, % (n=8)LPAH:Naphthalene 0.24 36Acenaphthylene nd ndAcenaphthene 2.5 8Fluorene 1.8 25Phenanthrene 15.9 19Anthracene 2.95 18HPAH:Fluoranthene 35.0 24Pyrene 30.5 23Benz(a)anthracene 38.0 23Chrysene 30.0 24Benzo(b)fluoranthene 12.7 14Benzo(k)fluoranthene 81.5 23Benzo(a)pyrene 41.7 28Dibenz(a,h)anthracene 38.5 36lndeno(1,2,3-cd)pyrene 22.7 28Benzo(g,h,i)perylene 22.6 30nd- not detectedLPAH-low molecular weight PAilsHPAH-high molecular weight PAHs139Another sediment sample (CD-i) obtained from Kitimat Arm near the Yacht Clubalso contained high concentration of PAHs, ranging from 0.5 ig/g of naphthalene to 53 igig of indeno(i,2,3-cd)pyrene.Assessment of possible in situ degradation of the mixed PAHs in this sediment wasattempted by measuring their concentrations, when the sediment was incubated with seawater from the same site at ambient temperature. No other nutrient, medium, or enrichedculture was added. Unfortunately any changes in the concentrations of the PAHs are smallcompared with the analytical variation resulting from the extraction, clean-up, and analysisprocedures. Thus no defmite conclusion can be reached regarding in situ degradation. Abetter method would be to measure the radioactivity of‘4C02 produced in the sedimentcontaining the added‘4C-labeled PAH, as described in Chapter 4. Although onlypreliminary studies are described above, the limited results indicate that the Kitimat Armenvironment provides an important source of microorganisms capable of degrading highmolecular weight PAHs, presumably because the microorganisms present in thisenvironment have been exposed to these PAHs.140Chapter 6. Preconcentration and Determination ofPolychiorinated Biphenyls (PCBs) and PAHsI. Simplex Optimization of a Capillary Gas Chromatography ElectronCapture Detection Systemand Solid Phase Extraction for the Determination of PCBs6.1 IntroductionPolychiorinated biphenyls (PCBs) are amongst the most widely distributedenvironmental pollutants (242-246). Their physico-chemical properties and ubiquitouspresence in the nature indicate that they are a probable threat to health (245-248).Because of their uses in transformers and capacitors, the concentration of PCBs in theenvironment varies with the extent of industrial discharge. Erickson (245) has listed PCBconcentrations in natural waters ranging from 30 pgIL to several hundred ngIL. In orderto measure PCBs at these trace levels in the environment, it is necessary to improve thesensitivity of analytical methods. To achieve this, improvement in instrumentation,optimization, and sample preconcentration are required. A great deal of effort has beenmade to the development of suitable instrumentation for trace analysis. Capillary gaschromatography (GC) with both mass spectrometry (MS) and electron capture detection(ECD) have become the methods of choice for the determination of PCBs (245, 246,249-25 1). However, sample clean-up and preconcentration continues to be a problem,and it has recently received much attention (252-255).A promising technique for sample handling is solid phase extraction (SPE). Tracecompounds of interest can be enriched on a suitable sorbent in order to isolate andpreconcentrate them prior to their separation and analysis. The sorbents are usuallypacked in a disposable cartridge or a short pre-column. Cartridges packed with polymer141resin XAD-2 (258, 259), Tenex (260), octadecylsilica (C 18) (261), and polyurethanefoam (262) have been used for the preconcentration of PCBs from water samples.Sample volumes are typically from 1 ml up to several liters (246, 252).In the present study, disposable glass Pasteur pipettes packed with C18 are usedfor the preconcentration/extraction of PCBs in water samples. Some problems commonlyassociated with the losses of PCBs during the sample handling are discussed, andsolutions to these problems are recommended.Another way to improve the sensitivity and separation of PCBs by GC is tosearch for the optimum GC operating conditions for the determination. Optimizationusing the traditional one-factor-at-a-time method is often time consuming, andsometimes fails to locate the maximum (263). Therefore, the simplex optimizationmethod (263-265) is used to delineate optimum conditions for the determination of PCBsby using capillary GC/ECD.6.2. Experimental6.2.1. InstrumentA Hewlett Packard 5890 gas chromatograph (GC) equipped with an electroncapture detector (ECD) was used throughout this work. A capillary GC column (DB5, 30m x 0.25 mm id., 0.25 urn film thickness; J & W) was used for the GC separation. AHewlett Packard 3393A integrator was used to record chromatograms and to integratepeak area and peak height.6.2.2. ReagentsA standard solution containing 10 PCB congeners (Table 6.1), representing 10isomeric groups was purchased from Supelco (PA, USA). Standard solutions of142commercial PCB mixtures (Aroclor 1221, 1242, and 1254) were also obtained fromSupelco. These solutions were diluted with an appropriate solvent when needed.The C18 packing material (particle size 10 jim) was purchased fromWaters/Millipore. All organic solvents used were of HPLC grade.Table 6.1. PCB congeners used in the present studiesNumber Name1 2-chlorobiphenyl2. 3,3’-dichlorobiphenyl3 2,4,5-trichiorobiphenyl4 2,2’,4,4’-tetrachlorobiphenyl5 2,3’4,5’6-pentachlorobiphenyl6 2,2’3 ,3’6,6’-hexachlorobiphenyl7 2,2’3,4,5,5’6-heptachlorobiphenyl8 2,2’,3,3’,4,4’,5,5’-octachlorobiphenyl9 2,2,3 ,3’,4,4’,5 ,5’,6-nonachlorobiphenyl10 2,2’,3,3’,4,4’,5 ,5’,6,6’-decachlorobiphenyl6.2.3. ProceduresGeneral procedures are described as follows. Disposable cartridges for solid phaseextraction were prepared by packing the appropriate amount of the C18 sorbent intoPasteur pipettes. A plug of glass wool was put near the outlet of the Pasteur pipette toretain the sorbent bed. Approximately 40 mg of C18 sorbent was added on the top of theglass wool in the pipette to prepare a bed of approximately 2-4 mm in height.143Approximately 1 ml of methanol followed by 1 ml of distilled water was applied to thecartridge before the cartridge was used to extract PCBs from water samples.Method one: Water samples (2-10 ml) were introduced to the top of thedisposable cartridge and allowed to drain freely through the cartridge. The cartridge wasthen dried by drawing air through it using an water aspirator. The analytes retained onthe cartridge were eluted by using toluene (100 111) and the eluent was collected in acalibrated glass sample vial. An aliquot (1-2 111) of the eluent was injected to the capillaryGC for analysis. Standard PCBs in toluene were used for calibration.Method two: To eliminate the loss of PCBs due to sorption on the glassware, thesample container was rinsed with toluene and this rinse solution was used to elute PCBsretained on the cartridge. The rest of procedures are the same as discussed in Methodone.Method three: A water sample in a container was put in an ice/water bath prior topassing through the cartridge, in order to reduce the possible loss of lower molecularweight PCBs through evaporation. The rest of procedures are the same as in Method two.The operating conditions of GC/ECD, established by using simplex optimizationand one-factor-at-a-time methods, are listed below. The injector temperature: 260 °C,splitless mode; the initial oven temperature: 104 OC; time remaining at the initialtemperature: 3.2 mm; the final temperature: 278 OC; the ramping rate from the initial tothe final temperature: 4.4 °C/min; time holding at the final temperature: 5 mm; thecapillary column head pressure: 21 psi; and the detector temperature: 290 °C.6.3. Results and Discussion6.3.1. Simplex Optimization144The operating conditions of GCIECD were optimized in order to achieve anefficient separation and to maximize sensitivity of detection for the PCBs. The simplexoptimization method (263-266) was used to optimize simultaneously the following sixexperimental variables: the injector temperature, the initial temperature of the GC oven,the time remained at the initial temperature, the final temperature, the linear ramping ratefrom the initial temperature to the final temperature, and the column head pressure. Thesum of mean resolution and mean peak intensity from the 10 PCBs (Table 6.1) waschosen as a response to be maximized.The total responses (the sum of mean resolution and mean peak intensity of 10PCBs) from each experiment as a function of variable value are displayed in Figures 6.1-6.3. Figure 6.1 shows the response at various initial temperature settings, which werechosen according to the rules governed by the simplex method. The initial temperaturehas been known to be very critical to the peak intensity and peak shape (267). It has beensuggested (267) that the optimum initial temperature should be approximately 10 °Cbelow the boiling point of the solvent used. The optimum initial oven temperature for thedetermination of PCBs in toluene solution, obtained by using the simplex method, was104 °C, which is 6 OC below the boiling point of the solvent used, toluene (110 OC). Thisresult is in agreement with those previously reported (267-269).The optimum initial time established was 2.5-3.5 mm as shown in Figure 6.2. Bymaintaining the column temperature slightly below the boiling point of the solvent forthis period of time, one is able to utilize the “solvent effect” (268, 269), resulting in animproved sensitivity.The optimum column head pressure was in the range 20-22 psi (Figure 6.3). Thecolumn head pressure is an indirect measure of the flow rate of carrier gas into thecapillary column.145100000090 0000 00 0o000000• •70 80 90 100 110 120Initial Temperature, CFigure 6.1. Responses obtained from 39 optimization experiments as a function of GCinitial temperature14610000000 090 e0 0e 00080 o0000070 o000 0601 2 3 4Initial Time, mmFigure 6.2. Responses obtained from 39 optimization experiments as a function of initialtime, held at the initial temperature14710000 000090 oO 000 0000o0C080 a0£2 00705 000eO60 • • • • •10 12 14 16 18 20 22 24Column Head Pressure, psiFigure 6.3. Responses obtained from 39 optimization experiments as a function of GCcolumn head pressure14870g60.aC*50 aLiiC.) ¼*< 40 I *a*30 • • i • •0 2 4 6 8 10Temperature Ramp, C/mmFigure 6.4. Resolution obtained from 39 optimization experiments plotted as a function oftemperature ramping rate149Varying the injector temperature between 255 and 270 °C had little effect on theresponse. However, peak intensities were reduced when the injector temperature wasbelow 255 °C, presumably because of incomplete sample transfer from the injector to thecolumn. Thus 255-270 °C should be chosen as the optimum injector temperature.The final oven temperature between 265 and 285 °C gave essentially constantresponse. No significant difference in either peak intensity or resolution was evident asthe final temperature was varied within this optimum range.The ramping rate from the initial to the final temperatures showed differenteffects on the resolution and on the sensitivity. The resolution decreases as the rampingrate increases (Figure 6.4); whereas the peak intensity gradually increases at the rampingrate varies from 2 to 8 °C/min. Therefore, different temperature rates may be chosenaccording to the application needs. In the present study, the optimum total responseinvolving both resolution and sensitivity was achieved at temperature rate 3-6 °Clmin.The 10 PCBs were all well separated from each other, and improving the resolution wasnot the primary objective. On the other hand, if the separation of a more complex PCBmixture is required, a lower ramping rate may be used in order to resolve closely elutingpeaks.6.3.2. Solid phase extractionPreliminary studies were carried out using Method one, a commonly used solidphase extraction procedure. An aliquot of 2.5 ml water sample containing 2-40 ng/ml of10 PCB congeners (Table 6.1) was allowed to pass through a disposable cartridgecontaining 40 mg of C18 sorbent. Recoveries were obtained by determining theconcentration of PCBs eluted from the cartridge with one of the commonly used solvents,ethylacetate, hexane, and toluene. Table 6.2 shows that the recovery of PCBs is not150complete, although this conventional method has been often used for thepreconcentration/extraction of several organics (252-254).When the cartridge was subsequently eluted using toluene, no PCBs was detectedin this second eluent, indicating that complete elution of PCBs was achieved by using100 tl toluene during the first elution. Therefore, the incomplete recovery of PCBs isprobably due to the insufficient retention of PCBs on the cartridge and/or the loss ofPCBs prior to the sample being introduced to the cartridge. Both possibilities wereinvestigated further as discussed below.The effect of pH of the water sample on the recovery of PCBs was studied, in anattempt to improve the retention of PCBs on the cartridge. It has been reported (270) thatthe recovery of some organics was affected by the acidity of water sample. In a series ofstudies, the pH of water samples was adjusted to 3, 6, 8, and 11, by using dilutehydrochloric acid and ammonium hydroxide solutions. No significant difference inrecovery of PCBs was found as the pH of the water sample varied from 3 to 8. Therecovery for the 10 PCBs was in the range of 30-80%, and this was reduced to 10-50%when the pH was increased to 11. Thus for the following studies, no pH adjustment onwater samples was needed, and the pH of the samples was generally in the range 4-8.Increasing the amount of the C18 sorbent from 40 mg to 300 mg showed noimprovement in recovery using Method one. The PCBs have a strong affinity for the C18sorbent used. It appears that breakthrough does not occur even in the narrow (severalmm) sorbent bed that was prepared from 40 mg of the C18 packing material. Verynarrow sorbent beds are now routinely used in a solid phase extraction technique basedon the use of an extraction disk that has been recently developed (270).Other possibilities for the low recovery of PCBs were examined. When 2.5 ml ofdistilled water and 10 il of PCB standard solution (in ethylacetate) were directly addedto the cartridge, drastic improvement in recovery over those shown in Table 6.2 wasachieved, especially for the large molecular weight PCBs. This indicated that the low151recovery obtained using Method one might have been due to the loss of PCBs before thewater sample was transferred to the cartridge for the preconcentration.Table 6.2. Recoveries of PCBs in water samples, obtained by using Method onePCBs Concentration Recovery (%)(ng/ml)Eluting Solvent:Ethylacetate Hexane Toluene1 40 75 50 892 40 56 64 723 4 48 64 604 4 37 51 495 4 33 43 436 4 34 45 457 2 31 35 358 2 35 34 389 2 35 34 3810 2 36 36 39Losses of PCBs due to the adsorption of PCBs onto the surface of the samplecontainer was suspected and studied in detail. Water samples (5 ml) spiked with knownamounts of PCBs were stored for approximately 30 mm in three separate containers:glass, polytetrafluoroethylene (PTFE), and stainless steel tubes, before the samples wereput to the disposable cartridge. The empty containers were each washed with toluene andthe wash solution was analyzed for PCBs. The percentage of PCBs found in the wash152solution as compared to the total amount of PCBs spiked were obtained and results aresummarized in Table 6.3. Up to 86% of the PCBs was found to be adsorbed on thesurface of the sample container. Thus, incomplete recovery noted using Method one isprobably due to analyte adsorption onto the glassware.Table 6.3. Percentage of PCBs adsorbed on the sample containersPCBs Adsorbed on containers (%)Glass vial PTFE vial Stainless steel vial1 - - -2 - - -3- 3 144- 9 335 12 23 516 20 28 567 34 59 578 44 74 579 46 82 5810 49 86 59153Results in Table 6.3 also suggest that PTFE adsorbs PCBs readily. Based on thisphenomenon, a sample preconcentration technique making use of a PTFE coil has beendeveloped and will be discussed later in Part II of this chapter.Most general SPE procedures, for example the EPA priority pollutants methods(271, 272), gave no special precautions regarding storage of water sample for PCBanalysis. They merely specify cold storage, sometimes in the dark or in amber bottles.The present results and the work of others (273-276) suggest that the sorption of PCBs tothe sample container can cause significant losses and appropriate care should beexercised. Pepe and Byrne (273) and Muidrew et at. (274) have noted the sorption of2,2’,4,4’,5,5’-hexachlorobiphenyl from water onto the container wall, causing significantlosses. Bellar and Lichtenberg (275) have suggested the use of formaldehyde to preservewater samples for PCB analysis. The addition of 20% (v/v) methanol was alsorecommended (276) to minimize the adsorption of polycyclic aromatic hydrocarbons(PAHs) onto container surfaces. In the present study, the PCBs that were adsorbed on thesample container were rinsed with toluene. The toluene rinse solution was then combinedto elute the PCBs from the cartridge. The adsorbed PCBs are therefore recovered alongwith the PCBs that were retained on the cartridge. As shown in Table 6.4, the recoveriesof PCBs are significantly improved (as compared to those in Table 6.2). Good recoveriesare achieved for the higher chlorinated PCBs, regardless of the time that the sample isstored in the glass container. Recoveries for the lower molecular weight PCBs, however,are not complete. As the sample storage time increases, the recovery of these PCBsdecreases. The lower recovery for these PCBs is probably due to their losses throughevaporation.154Table 6.4. Recoveries of PCBs obtained by using Method twoPCBs Recovery (%)After sample storage time (mm)2 20 2401 87 75 642 71 64 493 61 57 464 62 59 455 76 72 506 87 88 727 92 88 898 95 91 929 96 93 9710 96 94 98In the present studies, PCBs in ethylacetate were spiked to water samples atrelatively high concentrations, and PCBs have low water solubility. To prevent thepossible evaporation of PCBs along with the solvent (ethylacetate), an ice/water coldbath was used to store the water sample. Table 6.5 shows that complete recoveries wereobtained for all the 10 PCBs studied, when Method three was used. The combination oftwo precautions, storing the sample in a cold bath and rinsing the sample container withtoluene, eliminate the loss of PCBs, resulting in good recoveries for all the 10 PCBsstudied.155Table 6.5. Recoveries of PCBs obtained by using Method threePCBs Recovery (%)After sample storage time (mm)5 60* 1201 95 97±7 952 94 100±8 983 96 98±5 964 92 93±6 985 94 90±4 926 89 87±8 947 95 90±5 988 94 94±4 899 96 88±6 10310 89 92±5 100* mean ± standard deviation (n=4);Others are average values from duplicate experiments.Recoveries of PCBs in water samples at a range of concentrations were obtainedby using Method three, and results are summarized in Table 6.6. Quantitative recoveriesfor PCBs in concentration range 0.05-5 Hg/mi were obtained. A concentration factor of100 can be achieved by using 10 ml water sample and 0.1 ml toluene eluent.156Table 6.6. Recoveries of PCBs, at various concentrations spiked into water samples,obtained by using Method three.PCBs Cone. Recovery Cone. Recovery Cone. Recovery Conc. Recoveryng/ml % ng/ml % nglml % nglml %1 100 92 16.7 71 3.3 100 1 852 100 61 16.7 80 3.3 84 1 943 10 65 1.7 76 0.3 94 0.1 914 10 73 1.7 83 0.3 89 0.1 935 10 83 1.7 88 0.3 85 0.1 966 10 87 1.7 92 0.3 93 0.1 967 5 91 0.8 94 0.15 100 0.05 988 5 98 0.8 92 0.15 100 0.05 989 5 96 0.8 95 0.15 102 0.05 9510 5 96 0.8 95 0.15 99 0.05 96Relative standard deviations (RSD) from 5 replicate extractions of 10 ml watersample containing 0.1-2 ng/ml of PCBs were between 7 and 13%, and the averagerecoveries were between 85 and 98%.Similar procedures (Method three) were applied to the solid phaseextraction/preconcentration of PCB mixtures (Arochlor 1221, 1242, and 1254) spikedinto sea water samples, and good recoveries were also obtained, suggesting that thepresent method is useful for the preconcentration, extraction, and determination of PCBs.157II. In situ extractionlpreconcentration of PCBs and PAHs from aqueoussamples by using polytetrafluoroethylene (PTFE) tubingPrevious results shown in Table 6.3 indicate that PCBs are readily adsorbed on aPTFE surface. The loss of analyte by the adsorption can result in substantial error and isa nuisance in conventional sample analysis. However, this adsorption problem may beturned into an advantage in extracting/preconcentrating the analyte of interest. In thissection, the feasibility and application of using PTFE tubing for extracting/concentratingPCBs and PAHs from aqueous samples are examined. In situ extraction/preconcentrationof PCBs and PAHs are achieved by passage of aqueous samples through a PTFE tubing.The PCBs and PAHs present in the aqueous samples are removed from water andretained on the walls of the tubing while the water is allowed to flow through the tubing.The PCBs and PAHs that are trapped inside the tubing are then recovered by solventdesorption and the eluent is subsequently analyzed by using GC/ECD or GC/FID. Anumber of factors including the length of the tubing, the flow rate of the aqueous samplepassing through the tubing, and the volume and flow rate of the desorbing solvent, areinvestigated in order to maximize the recovery of trace analyte spiked into aqueoussamples.6.4. Apparatus and MethodsTwo systems for the preconcentration of PCBs and PAHs are schematicallyshown in Figure 6.5. In Scheme (a), a stream of nitrogen gas is used to drive the aqueoussample flowing through the PTFE tubing whereas a peristaltic pump is used in Scheme(b).In Scheme (a), a water sample is placed in a sample reservoir (150-mi separatoryfunnel) that is connected with the PTFE tubing. The sample flows through the tubing at aregulated flow rate, under an appropriate nitrogen pressure. The water that exits from the158tubing is collected in a separatory funnel containing 10 ml of extraction solvent(toluene). This toluene extract is analyzed by GC to check the efficiency of removal ofPCBs and PAHs from the aqueous sample (i.e. the retention of PCBs and PAHs on thetubing wall). A minimal volume of eluting solvent (toluene, 5-10 ml) was used to rinsethe sample reservoir, and was subsequently pushed through the PTFE tubing under thenitrogen pressure. The eluting solvent desorbs PCBs and PAHs from the tubing. Theanalytes were collected in a volumetric flask and were analyzed by using GC. Therecovery was obtained by comparing the amount of analyte eluted from the tubing withthe initial amount that was spiked into the aqueous sample.In Scheme (b), a peristahic pump was used to drive the sample through the PTFEtubing. The water that passed through the tubing was also collected and extracted withtoluene, to check the completeness of retention of analyte onto the tubing. In thedesorption process, an eluting solvent can be drawn back and forth with the aid of thepump so that the desorption efficiency can be improved. Four samples can be processedsimultaneously with the four-channel pump available. Also, the tubing can be directlyinserted into a water body at a particular depth and location for direct sampling. Nosample reservoir is necessary and therefore the possible adsorption of trace analyte ontothe reservoir is eliminated.15941*2j142NitrogenSampleTubing(a)Sample(b) Tubing PumpFigure 6.5. Schematic diagram of the preconcentration/extraction apparatus(I) collect the waste to check removal efficiency(2) collect eluent for analysis160Sea surface microlayer samples were obtained by using a similar method to thatdescribed by Harvey and Burzell (277), where a glass plate was immersed just under thewater surface to collect the surface microlayer (approximately 100 iim deep). Thesamples collected were processed immediately on shipboard by using the in situextraction/preconcentration technique as described above. A defined volume of a sample(200-500 ml) was allowed to pass through a 30-rn PTFE tubing (1.5 mm id.). Toluene(10 ml) was used to elute the adsorbed compounds. The toluene eluent was stored in arefrigerator for a week before being shipped to the laboratory for GC analysis. Both themicrolayer and water column samples from the same sampling location wereconcentrated and analyzed in the same manner.6.5. Results and DiscussionInitial experiments involving passing a 100 ml water sample containing 1.2-24ng/ml of PCBs through a 8-rn PTFE tubing (1.5 mm id.) show that PCBs are completelyremoved from the sample and retained on the PTFE tubing (Figure 6.6a). The PCBs thatare retained on the PTFE tubing can then be eluted with toluene (10 ml). Achromatogram from the analysis of the eluent is shown in Figure 6.6b. As compared tothe PCB standards (Figure 6.6c), quantitative recoveries for the last 7 PCBs are achieved,and a recovery of 50-80% is obtained for the first 3 PCBs. These results illustrate twopossible applications of the technique: (i) cleaning up of trace amounts of organicimpurities in water; and (ii) simultaneous preconcentrationlextraction of trace organicsfrom aqueous samples. Thus the removal efficiency, recovery, and the factors influencingthese are further studied in detail.1618(b)(a)Figure 6.6. GC/ECD traces from the analysis of (a) a water sample after passing througha S-rn PTFE tubing, (b) PCBs eluted from the PTFE tubing; and (c) astandard of 10 PCB5.(c) 53 427:i9 101626.5.1. RemovalTwo lengths of PTFE tubing, 8 m and 30 m, were used to study the efficiency ofthe removal of PCBs from the water sample. Table 6.7 shows the percentage of PCBsretained on the PTFE tubing, obtained by determining the concentration of PCBs in thewater sample after it passed through the tubing. Results are the average of duplicateexperiments. It is clear that most PCBs are completely removed from the water sample.Consequently, good recoveries for most of the 10 PCBs are achieved. It is expected that amore complete retention of PCBs was obtained on a longer tubing, as a longer time wasavailable for the water sample to be in contact with the PTFE tubing surface. However,further increase of the length of the tubing is not advisable because it would also increasethe time for the sample to run through the tubing, resulting in a longer analysis time.Therefore, a 30-m tubing was chosen as optimum in the present study.The flow rate of the sample through a given tubing also affects the percentage ofPCBs retained on the tubing, as it is illustrated in Table 6.8. As the flow rate increasesfrom 1 to 16.2 mI/mm, the percentage of PCBs retained on the tubing graduallydecreases, particularly for the first few PCBs. However, adequate retention of most of thePCBs were obtained when the flow rate was kept under 10 mI/mm. To ensure that allPCBs in the water sample are completely adsorbed on the tubing, a flow rate of 5 mI/mmwas chosen.163Table 6.7. Effect of the length of PTFE tubing on the removal and recovery of 10 PCBsfrom a water sample(Water sample flow rate: 5 mi/mm; Eluting solvent flow rate: 3 mllmin)PCBs Cone. Removal, % Recovery, %(nglml)8mtubing 30mtubing 8mtubing 3omtubing1 40 32 89 20 522 40 77 100 67 803 4 90 100 76 844 4 93 100 88 895 4 97 100 93 926 4 97 100 93 977 2 100 100 95 988 2 100 100 97 969 2 100 100 97 9410 2 100 100 98 97164Table 6.8. Percent of PCBs removed from water sample after the sample passed throughthe tubing at different flow ratesPCBs Removal (%)Flow rate (mi/mm):1 3 5 10* 13.8 16.21 97 94 85 70±13 55 482 100 97 100 90±3 82 663 100 100 99 89±3 85 694 92 96 91 93±6 87 695 97 96 95 90±8 92 826 96 96 97 94±6 91 877 98 97 99 95±6 96 888 97 98 99 94±8 96 899 97 98 99 94±8 95 8810 97 98 99 93±6 94 88* mean ± standard deviation, obtained from four replicate samples. Others were theaverage of duplicate experiments.6.5.2. Recovery and eluting parametersA number of eluting solvents including toluene, ethyl acetate, hexane, anddichioromethane were tested to desorb and to elute PCBs from the PTFE tubing. It wasfound that toluene was appropriate, because it eluted PCBs the most efficiently and it iscompatible for GC analysis (267). Therefore, toluene was used as eluting solvent.165Table 6.9 shows recoveries of PCBs when 2-10 ml of toluene was used to elutethe adsorbed PCBs from the 8-rn PTFE tubing. Increasing recoveries are generallyobtained as the volume of toluene used increases from 2 to 6 ml. No significantimprovement on PCB recoveries was observed when toluene volume was increasedfurther to 10 ml. Hence, 6-10 ml toluene was used for the present studies.Table 6.9. Recovery of PCBs when different volumes of toluene was used to elute PCBsfrom the 8 m tubing.PCBs Recovery (%)Toluene volume (ml):2 4 6 7 101 12 16 20 20 222 36 51 61 67 683 38 49 56 59 804 59 73 83 85 885 64 74 80 80 936 81 90 95 97 937 79 85 90 90 958 82 90 93 93 979 82 92 94 92 9710 84 93 96 96 98The flow rate of the eluting solvent also affects the recovery of PCBs. It wasfound that quantitative recoveries for most of the 10 PCBs were achieved using 10 ml oftoluene under a flow rate of 3 mi/mm (Table 6.10). When the eluting flow rate wasincreased to 10 mi/mm, the recovery was decreased due to an incomplete elution of allPCBs. However, a complete elution can be achieved by drawing the solvent to flow166through the PTFE tubing back and forth for two or three times, with the aid of theperistaltic pump (Figure 6.5b).Table 6.10. Recovery of PCBs obtained at different flow rates of the eluting solvent, 10 mlof toluenePCBs Recovery (%)Eluting flow rate (mi/mm):3 5 10 12.5 10*1 77±4 58±2 49±4 53 542 98±2 81±2 79±6 79 863 100±5 85±6 80±9 79 924 103±5 91±4 88±8 89 995 98±5 92±11 87±10 83 1016 100±6 92±8 88±10 87 1017 96±6 88±1 85±10 84 988 95±5 87±4 84±7 87 949 94±9 92±2 85±11 86 9810 89±8 91±1 83±10 85 93* The eluting solvent (10 ml toluene) was drawn, with the aid of a peristaltic pump, toflow back and forth in the tubing for three times.Mean standard deviation (n=4); others are the average from duplicate experiments.167Increasing the eluting temperature to 60 °C was also found to improve theefficiency of eluting the PCBs from the tubing. However, an ice/water bath was usedwhen collecting the eluent, in order to prevent the loss of PCBs from evaporation at theelevated temperature.6.5.3. PAHsSimilar procedures were also applied to the in situ preconcentration/extraction ofPAHs. Figure 6.7 shows the chromatograms obtained from a standard solution containing16 PAHs (a), the PAHs eluted from the tubing by using 10 ml toluene (b), and a watersample (spiked with the 16 PAHs) after passing through an 8-rn PTFE tubing (c).Complete removal of PAHs from the water sample was achieved by passing the samplethrough the tubing, and quantitative recoveries for most of the 16 PAHs are obtained.These preliminary results suggest that this technique can also be applied for thedetermination of PAHs after further optimization of the operating parameters.6.5.4. Effect of sample matrixTo evaluate if the present technique can be used for thepreconcentration/extraction of PCBs and PAHs from real samples, removal and recoveryof PCBs that are spiked into sea water were studied and results are summarized in Table6.11. It is clear that PCBs in sea water sample are also sufficiently retained onto andrecovered from the PTFE tubing. These results indicate that the technique can be used toextract trace amounts of organic compounds, such as PCBs and PAHs, directly from seawater samples. The volume of a sample from which PCBs or PAHs are extracted can becontrolled by regulating the flow rate and the time of drawing the sample into the tubingby using the peristaltic pump (Figure 6.5b).168Figure 6.7. GC/FJD traces from the analysis of (a) a standard Solution containing 16PAHs; (b) toluene eluent from the PTFE tubing after a water samplecontaining the 16 PAl-Is passed through the tubing; and (c) the water sampleafter passing through the tubing.C— C0.(a)L() C--aD(b)U,(c)169Table 6.11. Removal and recovery of PCBs from sea water samples that are spiked withPCB standards(Water sample flow rate: 5 mllmin; Eluting solvent flow rate: 3 mi/mm)PCBs Conc. Removal (%) Recovery (%)(ng/ml)8-rn tubing 30-rn tubing 8-rn tubing 30-rn tubing1 40 60 91 32 542 40 90 99 72 703 4 97 99 79 834 4 100 100 86 815 4 100 100 89 946 4 100 100 94 897 2 100 98 93 878 2 100 100 96 879 2 100 100 98 9710 2 100 100 95 971706.5.5. Application: In situ preconcentration/extraction of trace organics from sea surfacemicrolayer and sea waterFigure 6.8 shows typical chromatograms obtained from a sea surface microlayersample (a) and a water sample 10 cm below the sea surface (b). Both samples (200 mleach) were treated by using the in situ preconcentration/extraction technique onshipboard immediately after collection. Toluene (10 ml) was used to elute the adsorbedcompound and the toluene solution was analyzed by using GC/FID. It is shown in Figure6.8 that a higher concentration of organics is present in the microlayer sample than that inthe water sample 10 cm below the surface. Although the identity of the compoundsdetected by using GC/FID is not known, the present study demonstrates the potentialusefulness of the preconcentration/extraction technique developed here. The results alsoagree with the fmdings reported by others (278), who showed the enrichment of organiccompounds in the sea surface microlayer.Figure6.8.GCIFIDtracesfromtheanalysisof (a)aseasurfacemicrolayer sampleand(b)awatersample10cmbelowthesurface.172Chapter 7. Conclusions and Future WorkThe emphasis of this thesis is on the determination of PAHs and on studies of themicrobial degradation of PAHs. The Kitimat estuary, British Columbia was chosen as theenvironment for study, because it is contaminated by PAHs and little is known about thefate of PAHs in this environment. The results discussed in this thesis provide detailedinformation on the microbial degradation of phenanthrene and pyrene, leading to anincreased understanding of the fate of PAHs in the Kitimat environment.Mixed microbial cultures obtained from Kitimat Ann sea water and sedimentsamples can degrade phenanthrene, pyrene, and benzo(a)pyrene, by utilizing the formertwo as sole carbon source. These microorganisms are unique and potentially usefulbecause pyrene and benzo(a)pyrene are known to be very difficult to degrade. Previouslythere were only few reports on the microbial utilization of pyrene as sole carbon source.The present studies also provide valuable information on the possible degradation of otherPAHs by these microorganisms.The strain that is responsible for the degradation of phenanthrene was isolatedfrom a sea water culture, and is partially characterized by using the Gram test and Biologmetabolic analysis. It is now undergoing a series of microbiological studies throughcollaboration with Dr. J. Foght in the Department of Biological Sciences at the Universityof Alberta. Preliminary results indicate that the strain is possibly Pseudomonasfluorescensand it has some unique characteristics. Further studies are being undertaken to locate thegenes in the microorganism that are responsible for the degradation of phenanthrene.The degradation of phenanthrene, without the accumulation of any specificmetabolite, suggests that phenanthrene is probably mineralized to CO2 and H20. Thepotential mineralization of pyrene is demonstrated by measuring 14C02, one of themineralization products from the 14C-labeled pyrene. Up to 65% of the pyrene is found tobe mineralized after 38 days of incubation. A high efficiency of mineralization is173particularly important when bioremediation by using microorganisms is contemplated.Non-toxic end products of mineralization are also an advantage.The method described in the thesis for the determination of mineralization by usingradioactive tracer studies is very simple and useful. Because of the low background, themethod can be used to detect a low degree of mineralization. A drawback is that themethod does not provide chemical information on metabolites. Complementary to thismethod, analysis by using HPLC/UV absorption/fluorescence can provide quantitativeinformation on the changes of PAHs and their degradation metabolites. In the presentstudies, the contents of the whole culture flask were sampled for analysis: acetonitrile wasalso added to solubilize PAHs. This procedure has advantage in improved accuracy. Inmost previous studies, only an aliquot of an aqueous culture was usually sampled foranalysis and this inevitably introduces sampling error because many PAHs have very lowsolubility in aqueous solution.Considerable effort was made to identify the degradation metabolites becauseinformation on the metabolites is important for an understanding of the degradationprocesses. A technique based on the use of HPLC/absorption/fluorescence system and thesequential injection of the sample was shown to be effective for obtaining the absorptionand fluorescence spectra of a metabolite: conventional scanning spectrometers usually donot have high enough sensitivity to measure the trace amount of metabolite in the cultures.The combination of chromatography and spectrometry allows for an identffication of ametabolite as cis-4,5-dthydroxy-4,5-dihydropyrene, based on the identicalchromatographic retention time and absorption and fluorescence spectra of the metaboliteand the standard. The identification was confirmed by using GCIFIt) and GC/MS afteracetylation of the metabolite. The acetylation treatment is simple and is complete in anhour. The overall procedure is very useful for obtaining information from unknowncompounds in a sample at low concentration: lengthy preconcentration and isolationprocedures are eliminated.174It is important to relate the laboratory study to the natural environment; however,what happens in the laboratory does not always happen in the environment. In principle, atypical degradation metabolite can be used as an indicator for assessing the presence ofdegradation in the environment (in situ degradation). In the case of pyrene degradation,the typical metabolite, cis-4,5-dthydroxy-4,5-dihydropyrene accumulates in the initialstages of pyrene degradation and is further mineralized after the pyrene is completelydegraded. When pyrene is constantly available to the microorganisms, cis-4,5-dihydroxy-4,5-dihydropyrene should also be present in the culture. In the Kitimat Arm environment,there is a PAH input source from the aluminum smelter and pyrene is constantly availableto microorganisms. Therefore, searching for cis-4,5-dthydroxy-4,5-dihydropyrene inenvironmental samples is a rational approach for providing evidence on the in situdegradation of pyrene. The approach was successful and combined analytical techniquesincluding HPLC/absorptionlfluorescence, GC/FID, and GCIMS were used for theidentification and determination of cis-4,5-dihydroxy-4,5-dihydropyrene in sediment andpore water samples. The presence of cis-4,5-dihydroxy-4,5-dihydropyrene in these Kitimatsamples has significant implications, suggesting that indigenous microorganisms are ableto degrade pyrene in the natural environment. This is the first study showing the in situdegradation of any PAH in the natural environment. This approach of fmding in situmicrobial degradation evidence is effective. It involves intensive use of combinedanalytical techniques.Another approach is to perform degradation studies by using untreated samples.An example involving the mineralization of the 14C-labeled pyrene was described in thethesis. Up to 28% of the pyrene was mineralized in the sediment samples, supporting theresults on the in situ degradation of pyrene by these microorganisms.In order to gain some understanding of the degradation of pyrene in theenvironment, a number of factors were studied. The mixed cultures obtained are able todegrade pyrene at temperatures ranging from 2 to 26 °C. The degradation observed at a175temperature as low as 2 °C supports the contention that the degradation of pyrene undernatural conditions is possible. It was also observed that the degradation of phenanthreneand pyrene are enhanced when organic nutrients such as peptone and yeast extract areadded into the culture. This result has important implications regarding bioremediation. Itmay be possible in the future to degrade PAHs in the Alcan lagoons by using themicroorganisms obtained in this thesis work and by supplementing with appropriatenutrients. Mean lagoons contain very high concentration of PAHs, both in water andsediment. Reduction of PAH concentrations, by using microbial degradation in the lagooneffluents, before they are discharged into the Kitimat Arm is a promising approach forpollution control.The metabolite cis-4,5-dihydroxy-4,5-dihydropyrene is detected in both thecultures and the natural samples including pore water and sediments. It has highersolubility than the parent PAHs, therefore, it is easily accessible to aquatic organisms. Thiscompound is probably toxic. In the future, it will be important to study its effect onaquatic organisms such as crab since crab inhabitat the interface between water andsediment.The concentrations of non-polar organic contaminants in water samples are usuallyvery low because of their hydrophobicity. As a consequence, it is difficult to determinethese organics in water samples and preconcentration is necessary. It was observed thatthe surface of sampling containers strongly adsorbs trace organic compounds in watersamples, leading to the loss of trace organics during sample handling. Two methods forthe preconcentration of trace organic compounds have been developed, one making use ofhome-made cartridges containing C18 packing materials and the otherpolytetrafluoroethylene (PTFE) tubing. The method based on the use of PTFE tubing maybe used for simultaneous water sampling, preconcentration, and extraction of traceorganic compounds.176References1. 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