FACULTEIT LANDBOUWKUNDIGE EN TOEGEPASTE BIOLOGISCHE WETENSCHAPPEN

Academiejaar 2003 - 2004

PAH-BIODEGRADATION BY SPHINGOMONAS AND MYCOBACTERIUM : STUDY OF THEIR NATURAL ABUNDANCE, DIVERSITY AND NUTRIENT DEMANDS IN PAH-CONTAMINATED SOILS.

PAK-BIODEGRADATIE DOOR SPHINGOMONAS EN MYCOBACTERIUM : STUDIE VAN HUN NATUURLIJKE VERSPREIDING, DIVERSITEIT EN NUTRIËNT EISEN IN PAK-GECONTAMINEERDE BODEMS.

door

ir. Natalie Leys

Thesis submitted in fulfillment of the requirements for the degree of Doctor (Ph.D.) in Applied Biological Sciences

Proefschrift voorgedragen tot het bekomen van de graad van Doctor in de Toegepaste Biologische Wetenschappen

op gezag van Rector: prof. dr. apr. A. DE LEENHEER

Decaan: Promotoren: prof. dr. ir. H. VAN LANGENHOVE prof. dr. ir. W. VERSTRAETE prof. dr. ir. E. TOP dr. ir. D. SPRINGAEL dr. ir. L. BASTIAENS

ISBN 90-5989-016-7

Auteur en promotoren geven de toelating dit doctoraatswerk voor consultatie beschikbaar te stellen en delen ervan te kopiëren voor persoonlijk gebruik. Elk ander gebruik valt onder de beperkingen van het auteursrecht, in het bijzonder met betrekking tot de verplichting uitdrukkelijk de bron te vermelden bij het aanhalen van de resultaten van dit werk.

The author and the promoters give the authorization to consult and to copy parts of this work for personal use only. Every other use is subjected to the copyright laws. Permission to reproduce any material contained in this work should be obtained from the author.

Gent, 28 maart 2004

De auteur: ir. Natalie Leys

De promotoren:

Prof. dr. ir. Willy Verstraete, Dr. ir. Eva Top, Dr. ir. Dirk Springael, Dr. ir. Leen Bastiaens

" Research is what I'm doing when I don't know what I'm doing."

- Wernher von Braun –

Dankwoord

~ THE END ~

Cast

The Doctor…...... Natalie Leys

Partners in crime…………..……………………….…...... Karolien & Zita

Co-stars…...... Cindy, Barbara, David, Jan & Joke

Colleagues...... Annemie Ryngaert & the Vito crew

Assistants...... Students Els, Tine & Carlos

Supporters...... All my friends!

Boyfriend.…...... Joachim De Baer

Anti-stress ball……...... Yazoo the cat

Mom & Dad.…...... Erna Van Hool & Louis Leys

Sister & her family.…...... Isabella Leys & Ivan Eulaers & Cato

The in-laws...... Hilda Ringoot & Hugo De Baer Catherine De Baer & Jeroen Stuur

Dankwoord

Review comity…...... Prof. H. Van Langenhove, Prof. P. Sorgeloos, Prof. E. Vandamme, Dr. K. Smalla, Prof. M. Höfte, Prof. P. De Vos

Producer...... Prof. Willy Verstaete

Assistant Producer...... Prof. Eva Top

Director...... Dr. Dirk Springael

Assistant Director...... Dr. Leen Bastiaens

Logistics supervisor...... Dr. Ludo Diels

Production support.……….……………………………………The SCK crew

~

Thank you all very much !!!

~

That's all folks!

Summary

SUMMARY

Polycyclic aromatic hydrocarbons (PAHs) are major soil pollutants in many industrialized countries. Microbial degradation is considered to be the major route through which PAHs are removed from contaminated environments and therefore bioremediation is considered as a feasible remediation technology for cleaning PAH- contaminated soil. Mycobacterium and Sphingomonas strains using polycyclic aromatic hydrocarbons (PAHs) as sole source of carbon and energy could be essential members of such PAH-degrading bacterial communities, as they are often isolated during enrichments of PAH-degrading bacteria from such soil. Therefore, for future optimization of bioremediation process, it is of interest to study more in detail the distribution and diversity and specific nutrient requirements of Mycobacterium and Sphingomonas in PAH-polluted soil.

Four new culture-independent PCR-based detection methods targeting the 16S rRNA genes were developed to analyze PAH-degrading Mycobacterium and Sphingomonas communities in PAH-contaminated soils. Genus-specific primers were developed for PCR detection of either Sphingomonas species (Sphingo108f and Sphingo420r), or ‘fast-growing’ Mycobacterium species (Myco66f and Myco600r). The resulting amplicons were separated by Denaturing Gradient Gel Electrophoresis (DGGE) for generating Mycobacterium and Sphingomonas community fingerprints. Both Mycobacterium and Sphingomonas specific primer sets proved to be highly selective for the target group and single-band DGGE profiles were obtained for most strains tested. Strains belonging to the same species had identical DGGE fingerprints, and in most cases but not all, these fingerprints were typical for one species, allowing partial differentiation between species in a Mycobacterium or Sphingomonas population. Inoculated Sphingomonas and Mycobacterium strains could be detected at a cell concentration of 104 respectively 106 cells per gram of soil using the new primer set alone or 102 cells per gram of soil in a nested PCR approach in combination with eubacterial primers. In addition, 2 species specific primer sets were designed to detect bacteria related to Sphingomonas sp. EPA505 (EPAf and EPAr) and M. frederiksbergense (MYCOFf and MYCOFr). Using DNA extracts of a variety of inoculated PAH-contaminated soils, the EPA505 specific primer pair was able to

Summary detect EPA505 in concentrations as low as 102 cells per gram of soil. The MYCOF primer set could detect M. frederiksbergense in soil at a cell concentration of 104 cells per g soil via direct PCR and subsequent DNA-DNA hybridization of the PCR products or at a cell concentration of 102 cells per g soil via a nested PCR approach.

The new detection methods were used to rapidly asses the Mycobacterium and Sphingomonas population structure of several PAH-contaminated soils of diverse origin and different overall contamination profiles, pollution concentrations and chemical-physical soil characteristics. Using the Mycobacterium genus-specific detection method, fast-growing Mycobacterium species were detected in most uncontaminated soils and PAH-contaminated soils tested. By sequencing of cloned PCR products amplified from DNA from PAH-contaminated soil, well-known PAH- degrading species like M. frederiksbergense and M. austroafricanum were detected. However, in all PAH-contaminated soils bacteria were detected with 16S rRNA gene sequences related to the 16S rRNA gene of M. tusciae, a Mycobacterium species so far not reported in relation to biodegradation of PAHs. Using the species-specific detection method, M. frederiksbergense strains were detected in most PAH- contaminated soils, including soils in which no M. frederiksbergense strains were detected using the Mycobacterium genus-specific detection method. The new Sphingomonas specific PCR-DGGE method revealed the presence of Sphingomonas communities in all tested PAH-contaminated soils, with less diversity in soils containing highest phenanthrene concentrations. Sequence analysis of cloned PCR products revealed new 16S rRNA gene Sphingomonas sequences significantly different from sequences from known cultivated isolates. Sequences from environmental clones grouped phylogenetically with other environmental clone sequences available in data bases and possibly originated from several potential new species, not previously detected with culture-dependent detection techniques. In most of the tested PAH-contaminated soils, we detected also 16S rRNA gene fragments from Sphingomonas sp. EPA505 related strains.

By adding different inorganic supplements of nitrogen (N) and phosphorus (P) affecting the overall Carbon/Nitrogen/Phosphorus-ratio of soil, we investigated the impact of soil inorganic N and P nutrient conditions on PAH degradation by PAH- degrading Sphingomonas and Mycobacterium strains by means of soil slurry

Summary degradation tests. The general theoretical calculated C/N/P-ratio of 120/14/3 [expressed in mg] allowed rapid PAH metabolisation by Sphingomonas and Mycobacterium strains without limitation. In addition, PAH-degradation activity was not affected when circa 10 times lower concentrations of nitrogen and phosphorus were available, indicating that Sphingomonas and Mycobacterium strains are capable of metabolizing PAHs under low nutrient conditions. In addition, PAH-degradation was not affected by an excess of nitrogen and/or phosphorus unbalancing the C/N/P ratio in the soil. Supplements of nitrogen and phosphorus salts increased however the salinity of the soil slurry solutions and seriously limited or even completely blocked biodegradation.

The results presented in this thesis suggest an important role for Mycobacterium and Sphingomonas species in the PAH-degrading bacterial communities naturally colonizing PAH-contaminated soils with very different contamination profiles and different origin. Sphingomonas populations seem to dominate in soils contaminated with high concentrations of more bioavailable and more easily degradable PAHs such as phenanthrene, while Mycobacterium populations may be better adapted to flourish in soils enriched in less bioavailable higher molecular weight PAHs. In addition, the results and conclusions of the 4 year research presented in this thesis will enable us to improve bioremediation of PAH-contaminated soils by stimulating as efficiently as possible the key biodegrading organisms.

“Truth is ever to be found in the simplicity, and not in the multiplicity and confusion of things.”

- Sir Isaac Newton (1642-1727) -

Samenvatting

SAMENVATTING

Polycyclische aromatische koolwaterstoffen (PAK’s) zijn belangrijke chemicaliën die voorkomen in verontreinigde bodems in vele geïndustrialiseerde landen. Biologische sanering door middel van bacteriën, is een milieuvriendelijke technologie voor de zuivering van gronden vervuild met PAK’s. Microbiële afbraak is het belangrijkste proces dat zorgt voor de natuurlijke verwijdering van PAK’s in het milieu. Bacteriën van het Mycobacterium genus en het Sphingomonas genus maken mogelijk een essentieel onderdeel uit van de bacteriële gemeenschap die zorgt voor PAK-afbraak in de bodem. Bodem isolaten die in staat zijn om PAK’s te gebruiken als enige bron van koolstof en energie zijn in het verleden immers herhaaldelijke geïdentificeerd als Mycobacterium of Sphingomonas. Voor de optimalisatie van biologische PAK- afbraakprocessen, is het dan ook van belang om specifiek de verspreiding, diversiteit en specifieke voedingspatronen van deze groep van PAK-afbrekende Mycobacterium en Sphingomonas stammen verder in detail te bestuderen.

Vier nieuwe cultuuronafhankelijke detectie methoden werden ontwikkeld voor de analyse van PAK-afbrekende Mycobacterium en Sphingomonas gemeenschappen in gecontamineerde bodems. Twee sets van genus-specifieke primers homoloog aan het 16S rRNA gen werden ontwikkeld en gebruikt in een PCR-DGGE methode voor de simultane detectie van alle species van het Sphingomonas genus of van alle ‘snelgroeiende’ species van het Mycobacterium genus. Stammen die behoren tot hetzelfde species toonden werden gekenmerkt door identieke DGGE-profielen, en meestal was één bepaald DGGE-profiel ook kenmerkend voor één bepaald species. Deze PCR-DGGE techniek liet toe de verschillende Mycobacterium of Sphingomonas species in een natuurlijke gemeenschap te ontwarren. Met behulp van de nieuwe genus-specifieke primer sets kon men in een enkelvoudige PCR een minimale concentratie van 104 Sphingomonas cellen respectievelijk 106 Mycobacterium cellen per gram bodem detecteren, of circa 102 cellen per gram bodem via een ‘nested PCR’. Twee species-specifieke primer sets werden ontwikkeld voor de selectieve detectie van bacteriën verwant met Sphingomonas sp. stam EPA505 en M. frederiksbergense, twee species gespecialiseerd in PAK-afbraak. Met behulp van de nieuwe species- specifieke primer sets kon men in een enkelvoudige PCR een minimale concentratie

Samenvatting van 102 Sphingomonas sp. EPA505 cellen respectievelijk 104 M. frederiksbergense cellen per gram bodem detecteren, of circa 102 cellen per gram bodem via een ‘nested PCR’.

De nieuwe detectie methoden werden toegepast om snel de Mycobacterium en Sphingomonas populatie te karakteriseren van PAK-gecontamineerde bodems van diverse oorsprong en met verschillende contaminatieprofielen. In het merendeel van ongecontamineerde en PAK-gecontamineerde bodems ‘snelgroeiende’ Mycobacterium species gedetecteerd. In sommige PAK-gecontamineerde bodems werden stammen geïdentificeerd die sterk verwant waren aan wel gekende PAK- afbrekende species zoals M. frederiksbergense en M. austroafricanum. In alle PAK- gecontamineerde bodems werden 16S rRNA gen fragmenten gevonden met sterke gelijkenissen aan het 16S rRNA gen van M. tusciae, een Mycobacterium species dat tot op heden nog niet in verband werd gebracht met PAK-afbraak. De species- specifieke detectie methode onthulde de aanwezigheid van M. frederiksbergense stammen in bijna alle PAK-gecontamineerde bodems, zelfs in bodems waar de genus- specifieke methode geen Mycobacterium species had gedetecteerd. Daarnaast, werden ook in alle PAK-gecontamineerde bodems complexe Sphingomonas gemeenschappen gedetecteerd. In PAK-gecontamineerde bodems met de hoogste fenanthrene concentraties was de gedetecteerde Sphingomonas gemeenschap het minst gediversifieerd. Sequentie-analyse van gekloneerde PCR-fragmenten toonde een duidelijk onderscheid tussen de gedetecteerde Sphingomonas 16S rRNA genen en gekende genen van gecultiveerde Sphingomonas stammen. De gekloneerde PCR- fragmenten groepeerden met ander klonen sequenties beschikbaar in de elektronische databanken en behoren mogelijk toe aan een aantal nieuwe, tot op heden niet gecultiveerde, species. In de meeste PAK-gecontamineerde bodems werden via de species-specifieke methode ook Sphingomonas gedetecteerd die sterk verwant zijn aan Sphingomonas sp. EPA505.

De relatie tussen beschikbare concentraties van koolstof, stikstof en fosfor als voedingsbronnen in de bodem en het gedrag van PAK-afbrekende Sphingomonas en Mycobacterium stammen werd bestudeerd aan de hand van kleinschalige biodegradatietesten. Door toevoeging van anorganische stikstof (N) en fosfor (P) werden de concentraties en verhoudingen van koolstof/stikstof/fosfor (C/N/P-ratio)

Samenvatting van enkele natuurlijke bodems bijgestuurd. De PAK-afbraak door Sphingomonas en Mycobacterium stammen was snel en volledig onder condities die de algemene theoretisch bepaalde optimale C/N/P-ratio gelijk aan 120/14/3 (uitgedrukt in mg) benaderde. De afbraak werd zelfs niet gelimiteerd wanneer tien keer minder stikstof en fosfor beschikbaar was dan algemeen voorgeschreven als optimaal, wat er op duidt dat Sphingomonas en Mycobacterium in staat zijn om te groeien ten koste van PAK’s in omgevingen met beperkte voedingsbronnen. Meer nog, de PAK-afbraak door Sphingomonas en Mycobacterium werd niet gehinderd door een onevenwicht van de C/N/P-ratio door een overmaat aan stikstof of fosfor. Niettemin, werd er een sterke inhibitie of zelfs stop van de afbraak waargenomen wanneer door de toevoeging van stikstof en fosfor supplementen ook de zoutconcentratie te drastisch werd verhoogd.

De resultaten beschreven in deze thesis suggereren een brede verspreiding en een belangrijke rol voor Mycobacterium en Sphingomonas bacteriën in PAK-afbrekende microbiële gemeenschappen in gecontamineerde bodems. Sphingomonas populaties lijken te domineren in gecontamineerde bodems met hoge concentratie aan biobeschikbare PAK’s zoals fenantreen, terwijl Mycobacterium populaties zich beter lijken te handhaven in gecontamineerde bodems met lagere concentraties aan minder biobeschikbare hoogmoleculaire PAK’s. De resultaten en conclusies van het onderzoek beschreven in deze thesis, zal ons toelaten om in de toekomst biologische sanering van PAK-gecontamineerde bodems te verbeteren door selectief de belangrijkste organismen van de microbiële gemeenschap op te volgen en te stimuleren.

“There is no higher or lower knowledge, but one only, flowing out of experimentation.”

- Leonardo da Vinci (1452-1519) -

Contents

CONTENTS

INTRODUCTION ______1 PAH-BIODEGRADATION BY SPHINGOMONAS AND MYCOBACTERIUM :STUDY OF THEIR NATURAL ABUNDANCE, DIVERSITY AND NUTRIENT DEMANDS IN PAH- CONTAMINATED SOILS

CHAPTER 1 ______3 BACTERIAL PAH-BIODEGRADATION AND BIOREMEDIATION OF PAH-CONTAMINATED SOILS: A LITERATURE REVIEW

CHAPTER 2 ______61 OCCURRENCE AND DIVERSITY OF FAST-GROWING MYCOBACTERIUM SPECIES IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHs)

CHAPTER 3 ______83 MYCOBACTERIUM FREDERIKSBERGENSE, A MYCOBACTERIUM SPECIES SPECIALISED IN POLYCYCLIC AROMATIC HYDROCARBON (PAH) DEGRADATION, IS UBIQUITOUS IN PAH-CONTAMINATED SOILS

CHAPTER 4 ______97 OCCURRENCE AND PHYLOGENETIC DIVERSITY OF SPHINGOMONAS IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS)

CHAPTER 5 ______117 OCCURRENCE OF SPHINGOMONAS SP. EPA505 RELATED STRAINS IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS)

CHAPTER 6 ______129 INFLUENCE OF THE CARBON/NITROGEN/PHOSPHATE-RATIO ON PAH-DEGRADATION BY MYCOBACTERIUM AND SPHINGOMONAS STRAINS IN SOIL

CHAPTER 7 ______145 GENERAL DISCUSSION AND PERSPECTIVES

BIBLIOGRAPHY ______153

CURRICULUM VITAE ______181

“The task is, not so much to see what no one has yet seen; but to think what nobody has yet thought, about that which everybody sees.”

- Erwin Schrödinger (1887-1961) -

Introduction

INTRODUCTION

PAH-BIODEGRADATION BY SPHINGOMONAS AND MYCOBACTERIUM : STUDY OF THEIR NATURAL ABUNDANCE, DIVERSITY AND NUTRIENT DEMANDS IN PAH-CONTAMINATED SOILS

Polycyclic Aromatic Hydrocarbons (PAHs) are common pollutants of air, water and soil in many industrialized countries. Although PAHs are naturally present at low concentrations in the terrestrial environment, pollutions are mainly due to human activities. High concentrations of PAHs occur in contaminated soils at wood treating facilities, at sites formerly used to produce manufactured gas, petroleum processing plants and in river and harbor sludge. PAH-contamination is of environmental and public concern due to their toxic, mutagenic and carcinogenic properties.

Microbial degradation is considered to be the major route through which PAHs are removed from contaminated environments and therefore bioremediation is considered as a feasible remediation technology for cleaning PAH-contaminated soil. Currently, in situ and ex situ bioremediation techniques are, however, still considered ineffective for the efficient removal of PAHs from contaminated soil due to general low biodegradation rates obtained. Biodegradation is mainly hampered by the low bioavailability of the hydrophobic PAHs which strongly adsorb to organic soil material or dissolve in non-aqueous phase liquids. In addition, mostly the soil is basically treated as a ‘black box’ with the inability to control and direct the biological processes. Not much is known about the key organisms involved in the degradation processes and about the specific needs of the biocatalysts with respect to nutrition and environmental conditions for their optimal activity in the soil environment. Different research data indicate that certain groups of soil bacteria are specialized in colonization of PAH-contaminated environments and may play the main role in the biodegradation process. PAH-degrading isolates almost exclusively belong to the genera Sphingomonas and Mycobacterium, which seem to possess not only the necessary enzyme machinery for degradation of PAH compounds but also seem to make use of original strategies to enhance PAH bioavailability. However, current knowledge is mostly based on cultivation–based isolated strains. Not much is know

- 1 - Introduction about the in situ occurrence, distribution and PAH-degradation activity of Sphingomonas and Mycobacterium strains and their specific nutrient requirements to become stimulated.

The work presented in this thesis, aimed to elucidate the natural occurrence, diversity and nutrient demands of PAH-degradation by Mycobacterium and Sphingomonas in polluted soils. The first goal was the development of monitoring techniques to screen soils for the natural presence of bacteria belonging to the genera Sphingomonas and Mycobacterium and for monitoring their dynamics during bioremediation processes. The applicability of molecular techniques based on total soil DNA-extraction followed by specific 16S rRNA gene amplification by PCR and DGGE-analysis of the resulting amplicons was explored. The second aim was the determination of the impact of soil characteristics such as salinity, pH and nutrients available in the soil on the activity of PAH-degrading Sphingomonas spp. and Mycobacterium spp.. Different (in)organic supplements were added to the soil to change the overall C/N/P-ratio, the pH and/or the ionic strength of the soil and the degradation activity of added Mycobacterium and Sphingomonas spp. was followed.

The manuscript is divided in 7 chapters. Chapter 1 provides an overview of the current literature on microbial degradation of PAHs with emphasis on (i) the ecology of PAH-contaminated environments, (ii) the phylogeny of the bacteria and the catabolic systems involved in biodegradation, and (iii) the environmental parameters influencing microbial PAH-biodegradation activity in soil during bioremediation processes. Chapter 2, Chapter 3, Chapter 4 and Chapter 5 describe the development and testing of molecular monitoring techniques based on PCR and DGGE- fingerprinting to specifically detect Sphingomonas spp. and Mycobacterium spp. in soil. These new detection methods were used to study the occurrence and diversity of Sphingomonas and Mycobacterium communities in PAH-contaminated soil. In Chapter 6, we examined the different nutrition and environmental conditions to optimize the survival and activity of the PAH-degrading Sphingomonas spp. and Mycobacterium spp. in soil. Chapter 7 discusses the obtained results in the framework of the research objectives. Conclusions are drawn and perspectives for further research are presented.

- 2 - Literature Review

CHAPTER 1

BACTERIAL PAH-BIODEGRADATION AND BIOREMEDIATION OF PAH-CONTAMINATED SOILS: A LITERATURE REVIEW *†

* REDRAFTED AFTER: LEYS NATALIE, BASTIAENS LEEN, AND SPRINGAEL DIRK (IN PREPARATION)

BACTERIAL BIODEGRADATION OF PAHS IN CONTAMINATED HABITATS: A LITERATURE REVIEW, CURR.

ADV. APPL. MICROBIOL. BIOTECHNOL.

† REDRAFTED AFTER: LEYS NATALIE, BASTIAENS LEEN, AND SPRINGAEL DIRK (IN PREPARATION)

MICROBIAL BIOREMEDIATION OF PAH-CONTAMINATED SOIL: A LITERATURE REVIEW, CURR. ADV.

APPL. MICROBIOL. BIOTECHNOL.

INTRODUCTION

Polycyclic aromatic hydrocarbons (PAHs) are a group of highly stable aromatic organic compounds, consisting of benzene rings in linear, angular and cluster arrangements containing only carbon and hydrogen (Table 1-1) (Harvey 1991). The more rings, the more hydrophobic the compounds are, the higher their octanol-water partitioning coefficients (Kow), i.e., the lower their water solubility, and the higher their melting and boiling points (Table 1-1) (Ernst 1995). PAHs are of governmental concern as they pose a risk for ecosystems and public health because of their possible cytotoxic, teratogenic, mutagenic and carcinogenic properties (Table 1-1) (Enzminger 1987; Harvey 1991). Most unsubstituted PAHs with four or fewer rings, are relatively harmless for humans, but can be toxic to marine diatoms, gastropods, mussels, crustaceans, and fish. Unsubstituted PAHs with five or six rings exhibit a wide range of carcinogenic activity and can cause cancers in mammalian animals and humans. Substituted PAHs, i.e., methylated or nitrated PAHs, are even of more of concern for public health. The pathways for human exposure to PAHs are inhalation (active/passive smoking and inhaling of polluted air), ingestion (contaminated food and drinks) and skin adsorption (Menzie 1992).

- 3 - Chapter 1

TABLE 1-1 PHYSICAL AND BIOLOGICAL CHARACTERISTICS OF SOME PAHS (Ernst; Enzminger 1987; Harvey 1991)

Formula Melting Water solubility PAH Carcinogenity MW Point (°C) at 30°C (ppb) C H Naphthalene 10 8 82 31.700 NR 128,17 C H Acenaphtene 12 10 95 3.930 NR 152,21

C H Fluorene 13 10 116 1.980 NR 166,22 C H Phenanthrene 14 10 101 1.290 iiST, iiiA 178,24

C H Anthracene 14 10 218 73 ivST, ivA 178,24 C H Fluoranthene 16 10 110 260 iiST, ivA 202,26

C H Benzo(a)fluorene 17 12 190 NR NR 216,28

C H Pyrene 16 10 150 135 iST, ivA 202,26

C H Benzo(a)anthracene 18 12 161 14 iiST, iiA 228,29

C H Chrysene 18 12 256 2 iST, iA 228,28

C20H12 Benzo(b)fluoranthene 168 NR iiiST, iA 252,32

C H Benzo(k)fluoranthene 20 12 217 NR iiiST, iA 252,32

C H Benzo(a)pyrene 20 12 179 4 iiiST, iA 252,32

C H Dibenzo(ah)anthracene 22 14 270 NR iST, iA 278,33

C H Indenol(1,2,3-cd)pyrene 22 12 164 NR iiiST, iiA 276,34

C H Benzo(g,h,i)perylene 22 12 273 0,26 iiiST, iiiA 276,34

C H Coronene 24 12 442 NR NR 300,28

i – sufficiently proven. ii – limited proven. iii – insufficiently proven. iv – not proven. A – through animal testing. ST – through short term experiment. NR – not reported.

- 4 - Literature Review

PAHs are common pollutants of air, water and soil in industrialized countries (Edwards 1983). In the Flemish part of Belgium, for example, sites contaminated with PAHs are concentrated around the city of Antwerp and Ghent and counted in 2002 3908 sites with contaminated top soil and 295 sites with contaminated top soil and ground water (OVAM 2003). PAHs present in the atmosphere, surface soil and waters may originate from many different sources (Ernst 1995). Natural forest fires and volcanic activity release occasionally PAHs in the environment but the most prominent sources of PAH-contamination are related to anthropogenic processing of fossil fuels such as crude oil and coal. PAHs are worldwide distributed in the whole ecosphere due to increasing traffic emissions. The concentration of PAHs in urban atmosphere depends on the number and types of local emissions sources, temperature, weather and seasonal variations (Dann 2001), but in general concentrations in urban and industrial areas are 10-100 times higher than in more remote regions (Harvey 1991). PAHs enter water environments directly through precipitation of contaminated dust from the air, through the runoff of polluted ground and through pollution of rivers and lakes by municipal and industrial effluents (Harvey 1991; Ernst 1995). In addition, many river and harbor sediments are severely contaminated with PAHs. Maintenance of such water transport routes yields yearly millions of tonnes of heavily polluted sludge, which needs to be treated or dumped in confined landfills. PAH- contaminated soil is often found around former gas production plants, wood treatment plants where creosote was used, incineration plants, petroleum refining plants, and asphalt production plants.

Once introduced in the environment, PAHs accumulate in air, surface waters and sediments and soil and may persist for decades due to their hydrophobicity and recalcitrance. The fate of PAHs in the environment depends on many physical, chemical and biological interactions between the sorbate (PAHs), the sorbent (sediment/soil/tar), the solvent (NAPL/water) and the living organisms (Enzminger 1987; Heitzer 1993; Luthy 1997; Ramaswami et al. 1997; Gosh 2000). The relative importance of the individual processes is different for each PAH-compound and varies with the physicochemical properties of the site matrix and the environmental conditions. PAHs in the atmosphere are associated with airborne particles and are subject to various modes of chemical and photochemical degradation forming for example nitrogen and sulfur-containing heterocyclic compounds. Although the

- 5 - Chapter 1 concentration of PAHs in freshwater are usually low, PAHs dissolved in water can be ‘taken up’ by plants (Wild 1992), vertebrate fish and shellfish (Jackson 1994) and transferred in the food chain. Soils in particular are ‘sinks’ where PAHs tend to concentrate. The extractability and bioavailability of hydrocarbons in soils decreases with increasing soil hydrocarbon contact time, i.e., a process designated as ‘aging’ (Bauer et al. 1985; Weissenfels et al. 1992; Hatzinger et al. 1995; Macleod et al. 2000; Reid et al. 2000). The processes of hydrocarbon sequestration in soil are thought to be driven by (i) diffusion into soil micro pores, (ii) partitioning (ad- and absorption) into the soil organic matter (SOM), and (iii) accumulation in non-aqueous phase liquids (NAPL). Soils with higher organic carbon (OC) contents have a larger capacity to sequester hydrocarbons (Cornelissen et al. 1989; Weissenfels et al. 1992) and in soils with an OC content greater than 0.1%, partitioning into the SOM has been found to be the dominant sequestration process (Chiou et al. 1979; Means et al. 1980; Pignatello et al. 1996; Luthy 1997). Soil at manufactured gas plants are typically contaminated with dense NAPLs or coal tar particles, containing considerable amounts of PAHs (Efroymson 1994; Efroymson et al. 1995; Luthy 1997; Ramaswami et al. 1997). Mass transfer of PAHs between tar and the aqueous/solid phase is an extra process determining the fate of PAHs in such environment.

Unlike physico-chemical sequestration or transport, biodegradation is the only natural process for actual removal of PAHs from the environment (Sims et al. 1983; Cerniglia 1984; Cerniglia 1992). Biodegradation can be done by bacteria, fungi, plants or algae or synergetic consortia between these living organisms. Microbial degradation is considered as a major route through which PAHs are naturally removed from contaminated environments (Bossert et al. 1984; Cerniglia 1984; Enzminger 1987; Cerniglia 1992). Therefore, bioremediation is considered as an economically and ecologically beneficial remediation technology for the treatment of PAH polluted sites (Erickson et al. 1993; Wilson et al. 1993; Luthy et al. 1994; Würdemann et al. 1995).

- 6 - Literature Review

BACTERIAL BIODEGRADATION OF PAHS IN CONTAMINATED HABITATS

Ecology of PAH-degrading microbial communities in the environment

Microbial communities of contaminated soils can adapt to the contaminants and contribute to natural attenuation of pollutants such as PAHs. Different reports indicate that environmental stresses, including exposure to contaminants, lead to changes in microbial community structure through the selective enrichment of specific microorganisms that are more adapted to the new environment (Herbes et al. 1978; Spain et al. 1980; Spain et al. 1983; Wiggins et al. 1987; Bauer et al. 1988; Aeolin et al. 1989; Leahy et al. 1990; van der Meer et al. 1992; Tuhackova et al. 2001; Macleod et al. 2002). It has been shown that the community changes are controlled by the pollutant concentration, i.e., the number of specific degraders are higher and the degradation capabilities are more diverse in more contaminated soils (Spain et al. 1980; Spain et al. 1983; Wiggins et al. 1987; Aeolin et al. 1989; Leahy et al. 1990; Grosser et al. 1991; Grosser et al. 1995; Carmichael et al. 1997; Macleod et al. 2000; Tuhackova et al. 2001; Tuxen et al. 2002). Adaptation leads to reduced lag times and faster rates of PAH-mineralization and increases the extent of the degradation, i.e., a larger final proportion of PAHs was mineralized to CO2 (Bauer et al. 1988; Macleod et al. 2002). Microbial adaptation to PAHs in aquifers has been demonstrated by the mineralization of PAHs and contaminant-stimulated in situ bacterial growth (enrichment of PAH-degrading bacteria and enhanced numbers of Protozoa) in PAH- contaminated zones but not in adject uncontaminated zones (Madsen et al. 1991; Ghiorse et al. 1995).

Not much is currently known about the global taxonomic composition of the bacterial communities degrading PAHs in the environment and the interaction between the different members during the degradation process. There are only a limited number of reports on culture-based or culture-independent total community analysis of PAH- contaminated niches. The composition of the microbial community in PAH- contaminated habitats has mainly been assessed by the enrichment and study of PAH- degrading isolates.

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The identity of the isolated PAH-degrading bacteria depended more on the source environment and contamination profile than on the geographical origin of the source material. PAH-degrading bacteria have been isolated from many different anthropogenic contaminated environments (Bastiaens 1998; Bastiaens et al. 2000; Bicknell et al. 2001) as well as from undisturbed pristine environments (Heitkamp et al. 1989). PAH-degrading bacteria have been detected in biofilms on building stones (Ortega-Calvo et al. 1997), plant material (Juhasz et al. 2000a), top soil (road side soil) (Tuhackova et al. 2001; Johnsen et al. 2002), plant rhizosphere soil (Radwan et al. 1998; Daane et al. 2001), bulk soil (Song et al. 1986; Bouchez et al. 1995; Deziel et al. 1996; Dagher et al. 1997; Willumsen et al. 1997; Bastiaens 1998; Boonchan et al. 1998; Bastiaens et al. 2000; Juhasz et al. 2000a; Yuste et al. 2000; Rehmann et al. 2001; Baraniecki et al. 2002; Johnsen et al. 2002; Widada et al. 2002), subsurface soil and groundwater (Bicknell et al. 2001; Bakermans et al. 2002), fresh water and sediments rivers and lakes (Nortemann et al. 1986; Heitkamp et al. 1987; Cerniglia 1989; Dean-Ross et al. 2001), and, estuarine water and sediments from seas or oceans (Heitkamp et al. 1987; Heitkamp et al. 1988a; Cerniglia 1989; Geiselbrecht et al. 1998; Hedlund et al. 1999). Bacteria and fungi appear to be the dominant hydrocarbon degraders in soil environments while bacteria and yeast are the main degraders in aquatic ecosystems (Hanson et al. 1997). Via culture-independent analysis of 16S rRNA genes in soil-derived 13C-labeled DNA, Pseudomonas, Acinetobacter and Variovorax spp. were identified as the dominant active bacteria degrading 13C-labeled naphthalene in top soil (Padmanabhan et al. 2003). Oil- and PAH-utilizing rhizosphere isolates predominately belonged to the Arthrobacter genus (Radwan et al. 1998), Pseudomonas genus (Daane et al. 2001), Painibacillus genus (Daane et al. 2001) or Nocardia-Mycobacterium-Rhodococcus group (Daane et al. 2001). In moderated and heavily contaminated bulk soil, PAH-degrading isolates appeared to be mostly members of the Pseudomonas genus (Bouchez et al. 1995; Deziel et al. 1996; Dagher et al. 1997; Bastiaens 1998; Bastiaens et al. 2000), the Sphingomonas genus (Dagher et al. 1997; Mueller et al. 1997; Bastiaens 1998; Bastiaens et al. 2000; Ho et al. 2000; Baraniecki et al. 2002; Johnsen et al. 2002) and the Mycobacterium genus (Grosser et al. 1991; Bastiaens 1998; Bastiaens et al. 2000; Ho et al. 2000; Rehmann et al. 2001; Bogan et al. 2003), and in lesser extent members of the Rhodococcus

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(Walter et al. 1991; Bouchez et al. 1995; Tongpim et al. 1996) and Burkholderia (Mueller et al. 1997) genera. That Mycobacterium strains are ubiquitous in PAH- contaminated soils from very different origin was also confirmed by culture- independent 16S rRNA gene based community analysis (Cheung et al. 2001). Fresh water sediment PAH-degrading isolates were classified as a new species of the Polaromonas genus (Jeon et al. 2004). Obligate marine PAH-degrading isolates represented often new species and new genera of the γ-Proteobacteria such as Neptunomonas (Hedlund et al. 1999) and Cycloclasticus (Geiselbrecht et al. 1998; Kasai et al. 2002; Kasai et al. 2003).

In addition, a relationship can be observed between the complexity, the hydrophobicity and recalcitrance of PAH compounds and the bacteria using the compound. Most bacteria selected on 2-ring PAHs such as naphthalene belong to the Pseudomonas genus (Dunn et al. 1973; Simon et al. 1993; Kiyohara et al. 1994; Deziel et al. 1996; Bastiaens 1998; Bastiaens et al. 2000), Sphingomonas genus (Fredrickson et al. 1995; Yrjala et al. 1998; Davidson et al. 1999a; Davidson et al. 1999b) or exceptionally to the Rhodococcus genus (Larkin et al. 1999), Arthrobacter genus (Ortega-Calvo et al. 1994), Ralstonia genus (Widada et al. 2002) or Neptunomonas genus (Hedlund et al. 1999). Strains utilizing 3-ring PAHs such as phenanthrene belong mainly to the Sphingomonas genus (Mueller et al. 1990; Kästner et al. 1994; Balkwill et al. 1997; Mueller et al. 1997; Gibson 1999; Ho et al. 2000; Pinyakong et al. 2000; Cho et al. 2001; Johnsen et al. 2002; Borde et al. 2003), Pseudomonas genus (Menn et al. 1993; Bouchez et al. 1995; Mueller et al. 1997; Ortega-Calvo et al. 1997; Garcia-Junco et al. 2001a; Borde et al. 2003), Aeromonas genus (Kiyohara et al. 1976), Burkholderia genus (Mueller et al. 1997), or gram positive genera such as Nocardioides (Saito et al. 2000), Planococcus (Ortega-Calvo et al. 1997), Bacillus (Ortega-Calvo et al. 1997), and the actinomycte genera Arthrobacter (Schwartz et al. 2000), Nocardia (Ortega-Calvo et al. 1997), Rhodococcus (Walter et al. 1991; Bouchez et al. 1995; Tongpim et al. 1996; Dean- Ross et al. 2001), and Mycobacterium (Kleespies et al. 1996; Mueller et al. 1997; Churchill et al. 1999; Rehmann et al. 2001; Bogan et al. 2003). For the degradation of 4-ring PAHs such as pyrene a few gram negative strains belonging to the Proteobacteria genera Sphingomonas (Mueller et al. 1990; Weissenfels et al. 1991;

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Mueller et al. 1997; Kästner 1998), Burkholderia (Juhasz et al. 1997a), Pseudomonas (Boonchan et al. 1998) and Stenotrophomonas (Boonchan et al. 1998; Juhasz et al. 2000c) have been reported, but they mainly include members of the Corynebacterineae suborder, i.e., Gordonia (Kästner et al. 1994), Rhodococcus (Walter et al. 1991; Bouchez et al. 1995; Dean-Ross et al. 2001) and Mycobacterium (Grosser et al. 1991; Walter et al. 1991; Boldrin et al. 1993; Bouchez et al. 1995; Dean-Ross et al. 1996; Jimenez et al. 1996; Schneider et al. 1996; Thibault et al. 1996; Ho et al. 2000; Bogan et al. 2003; Gauthier et al. 2003). The apparent genus related preference for certain PAHs is probably due to increasing PAH-hydrophobicity with increasing number of aromatic rings. Some genera or species of PAH-degrading bacteria are more capable to interact with hydrophobic surface than others as also indicated by the effect of the isolation mode on the type of isolates. Using the same soils and the same PAHs compounds, different bacteria were isolated depending on the isolation procedure (Bastiaens 1998; Tang et al. 1998; Bastiaens et al. 2000; Friedrich et al. 2000; Grosser et al. 2000; Gauthier et al. 2003). Aqueous cultures using crystalline PAHs mainly led to the isolation of gram negative bacteria such as Pseudomonas and Sphingomonas species (Bastiaens 1998; Bastiaens et al. 2000). Two-liquid phase cultures selected for Mycobacterium, Bacillus, Microbacterium and Porphyrobacter strains (Gauthier et al. 2003). Enrichments using weakly sorbed PAHs (Amberlite beads) selected exclusively for Burkholderia species (Friedrich et al. 2000) whereas enrichments using strongly sorbed PAHs (hydrophobic Teflon membranes or polyacrylic beads) selected exclusively for Mycobacterium species (Bastiaens 1998; Bastiaens et al. 2000; Friedrich et al. 2000). Bacteria isolated on sorbed PAHs were more efficient in the degradation of the sorbed compound than species isolated from enrichment on non-sorbed PAHs (Tang et al. 1998), indicating that different PAH-degrading bacteria inhabiting the same soil may be adapted to different PAH-bioavailabilities. Analysis of microbial communities in sand-packed columns run with groundwater clearly showed compositional and functional difference in free-living and surface-associated bacteria that may reflect different roles of these distinct but interacting communities in the biodegradation of pollutants in aquifers (Lehman et al. 2002).

The dispersal of catabolic capacity via mobile genetic elements (MGE) and horizontal gene transfer (HGT) plays a major role in bacterial adaptation to environmental

- 10 - Literature Review stimuli, such as exposure to organic pollutants (Herrick et al. 1997; Stuart-Keil et al. 1998; Hohnstock et al. 2000; Park et al. 2003; Top et al. 2003; Wilson et al. 2003). Catabolic genes for PAH-degradation are often localized on conjugative plasmids and transposons (Park et al. 2003; Top et al. 2003; Nojiri et al. 2004). Dissimination of naphthalene-catabolic gene via conjugative plasmid transfer has been demonstrated in PAH-contaminated aquifers (Hohnstock et al. 2000). Conjugation frequency and natural HGT of PAH-catabolic plasmids was found to be stimulated by the exposure of the host to naphthalene (Hohnstock et al. 2000) and by the presences surfaces where conditions fostering stable, high-density cell-to-cell contact are more manifest (Park et al. 2003).

The distribution and diversity of the catabolic genes involved in PAH-biodegradation in the environment has only been studied very recently. Culture-independent PCR and RT-PCR techniques have indicated that naphthalene metabolism genes are ubiquiteous and actively in situ-transcribed in the indigenous community of coal-tar contaminated sites (Ghiorse et al. 1995; Wilson et al. 1999). The naphthalene metabolism genes were similar to those found on the NAH7 plasmid of P. putida strain G7 (nahAc and nahR genes) but significant sequence polymorphism of the gene and mRNA PCR-products indicated the presence of divergent homologs of the initial naphthalene dioxygenase alfpha subunit gene nahAc in the naphthalene-degrading bacterial population (Ghiorse et al. 1995; Wilson et al. 1999). Sequence comparisons revealed two major groups of nahAc homologs related to the naphthalene dioxygenase genes ndoB and dntAc, previously cloned from pDTG1 plasmid from P. putida NCIB 9816-4 and Burkholderia sp. strain DNT, repectively (Wilson et al. 1999). Culture- independent PCR and RT-PCR techniques have revealed much greater native naphthalene catabolic gene diversity than what was ever detected by culture-based approaches (Wilson et al. 1999). The ecological significance, relative distribution and transmission modes of the different naphthalene dioxygenase analogs was studied only rarely. One study on contaminated New Zealand soils indicated that the β- Proteobacterium phnAc allele, only found rarely in culture-based studies, may have a greater ecological significance in PAH-degrading communities than the extensively studied γ-Proteobacterium nah-like genotype (Laurie et al. 2000). A recent publication found that phnAc-like genes were mainly present in naphthalene- degrading isolates form a moderately contaminated hillside soil while nahAc-like

- 11 - Chapter 1 genes were found only among naphthalene-degrading isolates from an adjacent more heavily contaminated seep sediment (Wilson et al. 2003). Physiological characteristics of PAH-degrading bacteria

A large number of PAH-degrading microorganisms have been isolated from PAH- degrading environmental communities and characterized. Almost all PAH-degrading isolates are aerobic and able to use PAHs as sole source of carbon and energy. PAH- degrading bacteria of different genera seem to have adapted very specifically to the PAH-substrates they are using. Especially the genera Pseudomonas, Sphingomonas and Mycobacterium are well known for their degradation potential towards recalcitrant compounds including PAHs and have acquired diverse capabilities to inhabit a wide range of environments.

A typical adaptation of some Pseudomonas and Rhodococcus strains is the production of biosurfactants (e.g. glycolipids, sophorolipids, trehalose lipids, rhamnolipids, phospholipids and fatty acids) and bioemulsifiers (polysaccharides) which increase the aqueous solubility of PAHs (Deziel et al. 1996; Willumsen et al. 1997; Iwabuchi et al. 2002; Carcia-Junco et al. 2003). Some strains excrete extra-cellular surfactants but for most strains the surface activity is associated with the cell envelope (Willumsen et al. 1997).

Another adaptation is a strong adhesion to the hydrophobic PAH-substrate and the formation of biofilms on PAH crystals, on hydrophobic surfaces coated with sorbed PAHs and organic non-aqueous phase liquids with dissolved PAHs (Ortega-Calvo et al. 1994; Garcia-Junco et al. 2001b; Carcia-Junco et al. 2003). Biofilm formation can increase diffusion or dissolution of the PAHs. Some Pseudomonas and Sphingomonas strains enhance biofilm formation on PAH-substrates by the production of viscous extrapolysaccharides (EPS) (Pollock et al. 1999; Johnsen et al. 2000). The efficient PAH-degrading capacities of Sphingomonas and Mycobacterium strains are probably linked to these specific hydrophobic cell wall properties which control the interaction with and the membrane transport of hydrophobic compounds (Nohynek et al. 1995; Kawai 1999; Wick et al. 2001; Wick et al. 2002a). Both Sphingomonas and Mycobacterium strains have a very particular lipophilic outer cell wall layer. Glycosphingolipids replace the normal Gram negative lipopolysaccharides in the

- 12 - Literature Review outer membrane of Sphingomonas cells (Kawahara et al. 1999; Wiese et al. 1999; Yabuuchi et al. 1999) and Mycobacterium cells have an additional layer of mycolic acids (glycolipids) on top of the common Gram positive peptidoglycan cell wall (Sayler et al. 1994). Moreover, Mycobacterium cells respond to PAHs as growth substrates by changing the mycolic acid composition of their cell wall, i.e., they become more hydrophobic and more negatively charged (Wick et al. 2001; Wick et al. 2002a; Wick et al. 2002b; Wick et al. 2003b). While a more negative potential of cells would be expected to increase repulsive electrostatic interactions between cells and solids, a more hydrophobic cell wall might allow the bacterium to adhere better to hydrophobic surfaces such as Teflon and PAH crystals to form biofilms (van Loosdrecht et al. 1990a; van Loosdrecht et al. 1990b; Bastiaens 1998; Bastiaens et al. 2000; Wick et al. 2001; Wick et al. 2002a). PAH-degrading Pseudomonas, Sphingomonas and Mycobacterium strains have generally hydrophobic and negatively charged cell envelopes (Rijnaarts et al. 1993). The Gram positive Mycobacterium cells are in general more hydrophobic and more negatively charged than the Gram negative Pseudomonas and Sphingomonas cells (Rijnaarts et al. 1993).

Many PAH-degrading bacteria seem to use very specific substrate uptake mechanisms. There are strong indications that PAHs are metabolized by intracellular enzymes located in the periplasmic space or bound to membranes, indicating that transport of these compounds through outer membranes is a requisite for their metabolism (Kawai 1999). A novel pit dependant endocytosic macromolecule transport system for hydrophobic polymers has been identified in a Sphingomonas strain growing on alginate, a natural HMW hydrophobic polymer (Momma et al. 1999). Electron microscopy revealed dynamic changes in both cell surface and membrane structure and mouth-like pits which open en close depending on the presence or absence of the substrate. Similar systems may be used for the incorporation of PAHs. Also Mycobacterium species seem to be well adapted to the low available PAH concentrations as they make use of high-affinity uptake systems (Miyata et al. 2004) and can shift to maintenance metabolism during growth on poorly available sorbed PAHs (Wick et al. 2001; Wick et al. 2002a). A specific ‘interfacial’ uptake system has been suggested to enhance PAH-biodegradation by Arthrobacter and Rhodococcus cells absorbed at the interface of non-water-soluble non-degradable solvent containing PAHs (Ortega-Calvo et al. 1994).

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Some PAH-degrading strains are mobile and exhibit a chemotactic response towards PAHs (Pandey et al. 2002; Samanta et al. 2002). Pseudomonas putida G7, Pseudomonas sp. NCIB9816-4 and Pseudomonas putida RKJ1 are chemotactic attracted by naphthalene (Grimm et al. 1999; Samanta et al. 2000). For the first 2 strains, the naphthalene chemoreceptor, NahY, is encoded downstream of the naphthalene catabolic genes on the NAH7 plasmid and is co-transcribed with the catabolic genes (Grimm et al. 1999). In addition, also several Sphingomonas strains display phenotypic dimorphism and can adopt either a planktonic or sessile behavior in liquid media (Kovarova et al. 1998; Pollock et al. 1999; Johnsen et al. 2000). Sensing of environmental stimuli and genetic control over synthesis of the capsule are key events in alternating between these two phenotypes.

In addition, the simultaneous utilization of multiple substrates (mixed-substrate growth) has been shown to be a heterotrophic bacterial strategy under oligotrophic conditions (Kovariva et al. 1997; Kovarova et al. 1998) that might also be utilized by PAH-degrading bacteria. Simultaneous degradation of different PAHs has been reported several times (Guha et al. 1999). In ternary mixtures of PAHs the biodegradation rates of the more degradable and abundant compounds are reduced due to competitive inhibition, but enhanced biodegradation of the more recalcitrant PAHs occurs due to simultaneous biomass growth on multiple substrates (Guha et al. 1999). Some specialize PAH-degrading bacteria even utilize PAHs such as anthracene in the presences of much more easily degradable organic compounds such as glucose (Wick et al. 2003a). It could be a response to the heterogeneous composition of aromatic structures in the fossil organic matter in the habitat from which the strains were isolated (Romine et al. 1999a).

Taxonomy of PAH-degrading Mycobacterium and Sphingomonas strains

The genus of Mycobacterium [Lehmann and Neumann 1896] belongs to the phylum of Gram positive eubacteria with high G+C content (>55%), the class of Actinobacteridae, the order of Actinomycetales, the suborder of Corynebacterineae and the family of the Mycobacteriaceae (Tsukamura et al. 1977). The Mycobacterium

- 14 - Literature Review genus currently groups 100 different registered species and is represented by the type species Mycobacterium tuberculosis. The 16S and 23S rRNA gene based phylogenetic trees of the Mycobacterium genus show 2 major taxa : the ‘fast-growing Mycobacterium species’ form a coherent line of descent, distinct from the more recently evolved ‘slow-growing’ Mycobacterium species (Rogall et al. 1990; Stalh et al. 1990; Pitulle et al. 1992; Stone et al. 1995; Tortoli 2003). The ‘fast-growing Mycobacterium species’ are a group of Mycobacterium strains, mostly of environmental origin, that are, based on growth and biochemical characteristics and infectious properties (i.e. Mycobacterium species of Bio Safety Level 1, growth within 7 days), very different from the pathogenic and facultative pathogenic more slowly growing species including the overt pathogens such as M. avium, M. tuberculosis, M. leprae or M. ulcerans (i.e. Mycobacterium species of Bio safety level 2 & 3, growth after more than 7 days). The phylogenetic relatedness within the slow- and fast-growing corresponds in general with the classification based on traditional phenotypical analyses (Stalh et al. 1990). However, the current rRNA gene based phylogeny does not fully agree with previous numerical phenetic classification or Runyon classification based on pigmentation (Rogall et al. 1990; Pitulle et al. 1992). Both fast- and slow-growing species can be scotochromogenic (pigmented in light and dark), photochromogenic (pigmented in light) or non-chromogenic (not pigmented) (Tortoli 2003). So far, all PAH-biodegrading Mycobacterium isolates have been placed in the phylogenetic branch of the ‘fast-growing’ Mycobacterium species (Guerin et al. 1988; Briglia et al. 1994; Godvidaswami et al. 1995; Bastiaens 1998; Churchill et al. 1999; Poelarends et al. 1999; Yagi et al. 1999; Bastiaens et al. 2000; Schrader et al. 2000; Solano-Serena et al. 2000; Willumsen et al. 2001a). Most PAH-degrading mycobacteria are scotochromogenic and produce smooth round yellow colonies on solid media. PAH-degrading Mycobacterium isolates have been, based on 16S rRNA gene sequence, often assigned to the species M. frederiksbergense (Willumsen et al. 2001a), M. gilvum (Boldrin et al. 1993; Bastiaens et al. 2000; Vila et al. 2001; Gauthier et al. 2003), M. austroafricanum (Bogan et al. 2003), M. vanbaalenii (Heitkamp et al. 1988b; Godvidaswami et al. 1995; Wang et al. 1995; Khan et al. 2001; Moody et al. 2001; Khan et al. 2002), M. hodleri (Kleespies et al. 1996), M. flavescens (Dean-Ross et al. 1996), M. anthracenicum (Wang et al. unpublished) and M. chelonae (Kanaly et al. 2000a; Kanaly et al. 2000b; Kanaly et al. 2002).

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The Sphingomonas genus (Yabuuchi et al. 1999) was originally proposed to describe a group of bacterial strains isolated from human clinical specimens and hospital environments. The Sphingomonas genus belongs to the phylum of Proteobacteria, the class of α-Proteobacteria, the order of Sphingomonadales (i.e. the α-4 subgroup) and the non-photosynthetic family of the Sphingomonadaceae (Takeuchi et al. 1994; Kosako et al. 2000) and is represented by the type species Sphingomonas paucimobilis. The genus originally grouped 3 species but has quickly grown to 34 different registered species via reclassification of several old strains and addition of many new isolates (Balkwill et al. 1997; Yabuuchi et al. 1999; Hiraishi et al. 2000). 16S rRNA gene based phylogeny reflects classification of Sphingomonas based on chemotaxonomic characterization via polyamine patterns, polar lipid profiles and fatty acid composition (Busse et al. 1999). The phylogenetic tree of the Sphingomonadaceae family is complex as the heterogeneous braches belonging to the Sphingomonas genus are intermixed with branches of α-4 subgroup aerobic photosynthetic genera such as Porphyrobacter, Erythromicrobium, Erythrobacter and Sandaracinobacter. In 2001 proposed Takeuchi et al. to divide the heterogeneous Sphingomonas genus in the genus Sphingomonas sensu stricto and three new genera, Sphingobium, Novosphingobium and Sphingopyxis on the basis of phylogenetic and chemotaxonomic analyses (Takeuchi et al. 2001). However, in 2002 Yabuuchi et al. concluded that the genus Sphingomonas should remain undivided at this time, as none of the physiological and biochemical characteristics considered (including cellular lipids and fatty acid composition) provided evidence for the division of the current genus Sphingomonas (Yabuuchi et al. 2002). Based on 16S rRNA gene sequence, most of the cultured PAH-degrading Sphingomonas isolates (Mueller et al. 1990; Weissenfels et al. 1991; Kästner et al. 1994; Balkwill et al. 1997; Bastiaens 1998; Bastiaens et al. 2000) are close relatives of the S. yanoikuyae, S. herbicidivorans and S. chlorophenolica species type strains. These three species cluster phylogenetically in a group formally described as ‘the Sphingobium genus’ (Takeuchi et al. 2001). Over the last years, many new PAH- and pesticide-degrading strains and even new species such as S. xenophaga (Stolz et al. 2000), S. chungbukensis (Kim et al. 2000) or S. cloaca (Fujii et al. 2001) have been added to this ‘Sphingobium’ cluster and have emphasized the environmental importance of this group of Sphingomonas strains.

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Bacterial dioxygenases involved in biodegradation of PAHs

Bacterial PAH-degradation pathways

There are 3 types of microbial PAH degradation: mineralization, co-metabolic transformation and non-specific oxidation (Cerniglia 1984). Mineralization reactions will transform the substrate to molecules which can enter the central metabolism (acetate, pyruvate, citrate and methanol) and will lead to complete mineralization of the substrate to inorganic end products such as CO2 and H2O. These reactions will produce energy and carbon molecules which can be used for the production of cell material and reproduction (anabolic reactions). Bacteria that use PAHs as the sole source of carbon and energy, can grow and multiply on sole expense of the PAH compound. In co-metabolism, the PAH compound cannot support microbial growth but is modified and as such degraded when another growth-supporting substrate, mostly another PAH compound, is present. The energy gained from this transformation process is limited and can only support very low growth rates. In most cases, the co-metabolised PAH is not completely mineralized and metabolites may accumulate. Co-metabolism is considered as an important mechanism to degrade PAHs with higher molecular weigths in mixtures of PAHs (Alexander 1980; Keck 1989). Non-specific oxidation of PAHs is based on the activity of highly non-specific enzymes such as the extracellular lignine peroxidase produced by white rot fungus Phanerochaete chrysosponium (Bogan et al. 1996).

Chemically, the mineralization of PAHs is a chain of oxidation and reduction reactions. The most common used final electron-acceptor in PAH-degrading bacteria is molecular oxygen. Anaerobic PAH-biodegradation using nitrate or sulfate as final electron acceptor has been recently described but the enzymes and genes involved are not yet identified (Sharak Genthner et al. 1997; Zhang et al. 1997; Rockne et al. 2001; Chang et al. 2002; Eriksson et al. 2003). The complete biochemical pathways for aerobic microbial biodegradation of aromatic compounds including PAHs have been well described (Cerniglia 1984; Cerniglia 1989; Cerniglia 1992; Sutherland et al. 1995; Kanaly et al. 2000b) and can be consulted at the website of

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Biocatalys/biodegradation Database of the University of Minnesota (http://umbbd.ahc.umn.edu). Initial PAH-hydroxylating dioxygenases

Enzymatically, the initial step in the aerobic catabolism of aromatic rings by bacteria involves an oxygenase that catalyzes the ring oxidation producing vicinal cis- dihydrodiols. Therefore bacteria produce monooxygenases (incorporate 1 oxygen atom) and dioxygenases (incorporate 2 oxygen atoms) while fungi and mammals only produce monooxygenases. Initial attack of PAHs by bacteria involves substrate specific dioxygenases. Bacterial aromatic ring-hydroxylating dioxygenases are multicomponent enzyme systems that consist of a reductase (flavoprotein), a ferredoxin, and a terminal dioxygenase (Figure 1-1). The reductase and the ferredoxin transport electrons to the terminal dioxygenase which catalyzes the reaction. In the course of the reaction two oxygen atoms, two electrons and two protons are consumed. The terminal dioxygenase, an iron sulphur protein (ISP), is composed of a large (α) and a small (β) subunit. The α-subunit (ISPα) is the catalytic component and contains 2 conserved regions: the Rieske [2Fe-2S] center and the mononuclear iron domain, which are involved in the consecutive electron transfer to the dioxygen molecule. Both α- and β-subunits of the ISP are necessary for function and in determining the substrate specificity of the dioxygenase. The initial ring oxidation with substrate specific dioxygenases is considered the rate-limiting step in the biodegradation of PAHs.

PAHs

dihydrodiol

FIGURE 1-1 ELECTRON TRANSPORT BETWEEN THE SUBUNITS OF A INITIAL PAH-DIOXYGENASE

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The aromatic ring-hydroxylating dioxygenases responsible for the first step in the aerobic oxidation of PAHs are substrate and bacterium variable. Naphthalene dioxygenase have mainly been isolated from γ-Proteobacteria, i.e., from a few strains from genera such as Neptunomonas (Hedlund et al. 1999), Cycloclasticus (Geiselbrecht et al. 1998; Kasai et al. 2003; Hedlund et al. unpublished), Pseudoalteromonas (Hedlund et al. unpublished) or Marinobacter (Hedlund et al. unpublished), but mainly from the Pseudomonas genus (Schell 1986; Kurkela et al. 1988; Haigler et al. 1990; Kelley et al. 1990; Herrick et al. 1993; Simon et al. 1993; Dagher et al. 1997; Kosheleva et al. 1997; Geiselbrecht et al. 1998; Bosch et al. 1999b; Hamann et al. 1999; Mordukhova et al. 2000; Park et al. 2000; Ferrero et al. 2002; Park et al. 2002; Olivera et al. 2003) (Table 1-2). The operons coding for naphthalene dioxygenase in Pseudomonas strains are (i) nahAaAbAcAd genes (Schell 1986; Herrick et al. 1993; Simon et al. 1993; Bosch et al. 1999b; Ferrero et al. 2002; Park et al. 2002; Olivera et al. 2003), (ii) ndoAaAbAcAd genes (Kurkela et al. 1988; Simon et al. 1993; Yang et al. 1994; Pellizari et al. 1996; Hamann et al. 1999), (iii) pahAaAbAcAd genes (Kiyohara et al. 1994; Takizawa et al. 1994), and (iv) doxAaAbAcAd genes (Denome et al. 1993). In general, the geneAa encodes the reductase, the geneAb for the ferredoxin, the geneAc for the large α-subunit of the ISP and the geneAd for the small β-subunit of the ISP. Naphthalene dioxygenase genes have recently also been sequenced from β-Proteobacteria such as Commamonas (Moser et al. 2001; Jeon et al. 2003), Ralstonia (nagAaAbAcAd genes) (Fuenmayor et al. 1998; Zhou et al. 2001; Widada et al. 2002), Polaromonas (Jeon et al. 2003), Burkholderia (phnAaAbAcAd genes) (Wilson et al. 2003), Herbaspirillum (Wilson et al. 2003) or Bordetella (Parkhill et al. 2003a), from α-Proteobacteria such as Sphingomonas (bph genes or nsaAaAbAcAd genes) (Romine et al. 1999b; Conradt et al., unpublished), and from some Gram positive bacteria such as Mycobacterium (nid genes) (Khan et al. 2001) or Rhodococcus (narAaAbAcAd genes or bpfAaAbAcAd genes) (Larkin et al. 1999; Treadway et al. 1999; Andreoni et al. 2000; Kulakov et al, unpublished). Related genes have also been found in the genome of Mesorhizobium (Kaneko et al. 2000) or Agrobacterium (Wood et al. 2001) strains or even in an Archae bacterium Thermoplasma (Ruepp et al. 2000). Naphthalene dioxygenases are very versatile enzymes that can catalyze not only the degradation of naphthalene but

- 19 - Chapter 1 catalyze also many dioxygenation and monooxygenation reactions with other aromatic hydrocarbons, including substituted aromatic hydrocarbons and heterocyclic aromatic hydrocarbons (Resnick et al. 1996). The naphthalene dioxygenase of P. putida NCIB 9816/11, also catalyzes the fluorene monooxygenation or dioxygenation. The naphthalene degradation enzymes of other Pseudomonas strains are also involved in the transformation of phenanthrene and anthracene (Menn et al. 1993; Sanseverino et al. 1993; Yang et al. 1994).

Genes encoding specific dioxygenases involved in the initial attack of fluorene have to our knowledge only been sequenced from one bacterium, i.e., the Actinomycetes Terrabacter (fln genes) (Habe et al. unpublished) (Table 1-2). Two different dioxygenation fluorene degradative routes which support growth by production of central metabolites have been described (Cerniglia 1992; Grifoll 1992; Boldrin et al. 1993; Monna et al. 1993; Resnick et al. 1996; Casellas et al. 1997; Wattiau et al. 2001; van Herwijnen et al. 2003b). For most strains, initial attack occurs by a monooxygenation at the C-9 of fluorene followed by an angular-carbon dioxygenation.

Specific phenanthrene dioxygenase genes have mainly been sequenced from β- Proteobacteria such as Commamonas (Goyal et al. 1996), Burkholderia (phn genes) (Laurie et al. 1999a; Laurie et al. 1999b) and Alcaligenes (Kiyohara et al. unpublished). The genes and enzymes involved in phenanthrene degradation pathway have recently also been characterized from 2 α-Proteobacteria Sphingomonas strains (adhAaAbAcAd genes) (Pinyakong et al. 2003; Iwabuchi et al. unpublished) and 4 Actinomycetes of the genera Nocardioides (phdAaAbAcAd genes) (Iwabuchi et al. 1998; Saito et al. 2000) or Mycobacterium (pdoAaAbAcAd genes or nidAaAbAcAd genes) (Krivobok et al. 2003). In the phenanthrene and pyrene degrading Mycobacterium sp. 6PY1, two operons encoding a phenanthrene dioxygenase enzyme complex have been sequenced (pdo1AaAbAcAd and pdo2AaAbAcAd genes) (Krivobok et al. 2003). In contrast to the strict selective phenanthrene dioxygenases Phd and Pdo2, the Nid and Pdo1 dioxygenase can also catalyze the dihydroxylation of pyrene. Phenanthrene dioxygenases initiate phenanthrene degradation mostly by an initial dioxygenation at the 3,4-position (Cerniglia 1984). It has been demonstrated that in several Pseudomonas strains naphthalene and phenanthrene share a common

- 20 - Literature Review upper metabolic pathway, i.e., a common set of enzymes is responsible for the conversion of phenanthrene to 1-hydroxy-2-naphthoic acid as well as that of naphthalene to salicylic acid (Kiyohara et al. 1994; Yang et al. 1994) (Table 1-2). The 1-hydroxy-2-naphthoic acid is oxidized to 1,2-dihydroxynaphthalene, which is further metabolized to salicylate via the naphthalene pathway (Evans et al. 1965; Balashova et al. 2001). In β-Proteobacteria, 1-hydroxy-2-naphthoic acid undergoes ring-cleavage and is further metabolized via o-phthalate and protocatechuate (Kiyohara et al. 1976; Iwabuchi et al. 1998; Pinyakong et al. 2000; Shuttleworth et al. 2000). Recent studies, have shown that for phenanthrene also dioxygenation at the 1,2-position followed by meta-cleavage is possible (Jerina et al. 1976; Pinyakong et al. 2000). In addition, the metabolism of phenanthrene by Streptomyces flavovirens and the marine Cyanobacterium Agmenellum quadruplicatum PR-6 is more similar to that reported in mammalian and fungal enzyme systems than those catalyzed by bacteria. Both oxidize phenanthrene to phenanthrene trans-9,10-dihydrodiol via a monooxygenase- epoxide hydrolase-catalyzed reaction rather than by a dioxygenase.

Not much is known about the genetics of specific anthracene dioxygenases. The fluoranthene degradation genes of the α-Proteobacterium Sphingomonas paucimobilis strain TNE12 have been localized on a 240kb plasmid (Shuttleworth et al. 2000), but have not yet been sequenced.

Only a few bacterial strains have been identified that can completely metabolize the 4- ring PAH pyrene to CO2, and most of them belong to the Mycobacterium genus. Pyrene degradation enzymes and genes have been identified in pyrene degrading Mycobacterium strains M. vanbaalenii PYR-1 (nid genes), Mycobacterium sp. PAH2.135 (RIGII-135) (nid like genes), M. flavescens PYR-GCK (ATCC 700033) (nid like genes), M. gilvum BB1 (DSM 9487) (nid like genes), M. frederiksbergense FAn9 (DSM 44346T) (nid like genes) and Mycobacterium sp. 6PY1 (pdo2 genes). One copy of nidA and nidB like genes were detected in strain PAH2.135, while multiple copies of each gene were detected in the strains PYR-1, BB1, PYR-GCK and FAn9 (Brenza et al. 2003). The different copies could be essentially identical copies or different homologous genes coding for different ring-hydroxylating dioxygenases within the same strain, similar to phenanthrene and pyrene dioxygenase Pdo1 and phenanthrene specific dioxygenases Pdo2 that are co-expressed in Mycobacterium sp.

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6PY1 (Krivobok et al. 2003). As mentioned before, for some Mycobacterium strains such as M. vanbaalenii PYR1 (Nid) (Khan et al. 2001) and Mycobacterium sp. 6PY1 (Pdo1) (Krivobok et al. 2003) it has been shown that the pyrene dioxygenases can catalyze dioxygenation of both phenanthrene and pyrene (Table 1-2). Furthermore, all Mycobacterium strains described so far to grow on pyrene also metabolize phenanthrene. In addition, in many Mycobacterium strains, pyrene degradation is induced by phenanthrene (Molina et al. 1999). These findings suggest that the same enzyme systems are involved in the catabolism of phenanthrene and pyrene in Mycobacterium strains, which is consistent with the current knowledge on the catabolic pathways for these 2 PAHs (Krivobok et al. 2003). The biosynthesis of the catabolic enzymes responsible for pyrene degradation in Mycobacterium strains seems to be under strict metabolic control, i.e., production is only induced by the presence of PAHs or their pathway intermediates (Krivobok et al. 2003). Pyrene degradation in Mycobacterium cells proceeds preferentially through the dioxygenation initiated Kivonara pathway (Heitkamp et al. 1988b; Cerniglia 1992; Boldrin et al. 1993; Krivobok et al. 2003).

PAHs with more than 4 rings also called high-molecular-weight PAHs (HMW-PAHs) are very recalcitrant or even persistent. So far, no bacteria have been isolated to use these PAHs as source of carbon and energy. Such high-molecular-weight PAHs (HMW-PAHs) are removed from the environment trough cometabolism, using other low-molecular-weight PAHs (LMW-PAHs) as growth substrates. Cometabolic degradation of PAHs with more than 4 rings had been reported for a Mycobacterium (Schneider et al. 1996), a Burkholderia (Juhasz et al. 1997b), and some methanotrophs. Cometabolisation of the potent carcinogen benzo(a)pyrene is known to proceed through substituted pyrene intermediates (Schneider et al. 1996) and depends on the presence of pyrene as cosubstrate (Boonchan et al. 2000; Juhasz et al. 2002).

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TABLE 1-2 GENES CODING FOR INITIAL PAH-HYDROXYLATING DIOXYGENASES IN BACTERIA

Catabolic genes‡ Host strains Location* Reference Naphthalene 1,2-dioxygenase nahAaAbAcAd Pseudomonas putida G7 pNAH7 C (Schell 1986; Harayama et al. 1989) Pseudomonas putida pNPL1 (Boronin et al. 1989) Pseudomonas putida PaW736 (NCIB 9816-4) pDTG1 C (Simon et al. 1993; Park et al. 2002) Pseudomonas putida 3IIIA2, 5IIANH, 5IIIASal NR (Ferrero et al. 2002) Pseudomonas putida 2IDINH, 3IA2NH, PR1MN1 NR (Ferrero et al. 2002) Pseudomonas putida Cg1 pCg1 C (Park et al. 2003) Pseudomonas stutzeri AN10, AN11 NR (Bosch et al. 1999a) Pseudomonas stutzeri B2SMN1, S1MN3, LSMN3 NR (Ferrero et al. 2002) Pseudomonas stutzeri STMN3, ST27MN3, LSMN2 NR (Ferrero et al. 2002) Pseudomonas stutzeri ST27MN2 NR (Ferrero et al. 2002) Pseudomonas stutzeri 63, 67, 85 NR (Olivera et al. 2003) Pseudomonas balearica LS402, SP401, SP1402, LS401 NR (Ferrero et al. 2002) Pseudomonas fluorescens A24, A88, I-16 NR (Izmalkova et al. unpublished) Pseudomonas fluorescens LP6a pLP6a (McFarlane et al. unpublished) Pseudomonas fluorescens NR (Min et al. unpublished) Pseudomonas aeruginosa SCD-1, S1-1 NR (Duncan et al. unpublished) Pseudomonas sp. PR3MN2, 8IDINH, LSMN7, 19IIDNH NR (Ferrero et al. 2002) Pseudomonas sp. ND6 pND6-1 (Li et al. unpublished) Pseudomonas sp. 5N1-1, 4N4-1, 4N1-3, 4N1-2, 4N1-1 NR (Bosch et al, unpublished) Pseudomonas sp. SOD-3, SCD-3a, SCD-14b NR (Duncan et al. unpublished) Burkholderia sp. SOD-5b, S1-17 NR (Duncan et al. unpublished) Cycloclasticus pugetii PS-1 NR (Geiselbrecht et al. 1998) Cycloclasticus sp. W NR (Geiselbrecht et al. 1998) Neptunomonas napthovorans NAG-2N-126, NAG-2N-113 NR (Hedlund et al. 1999) Mesorhizobium loti MAFF303099 NR (Kaneko et al. 2000) Agrobacterium tumefaciens C58 NR (Wood et al. 2001) Streptomyces coelicolor A3(2) NR (Bentley et al. 2002) Comamonas testosteroni GZ42 NR (Jeon et al. 2003) Polaromonas napthalenivorans CJ2 NR (Jeon et al. 2003) Marinobacter sp. NCE312 NR (Hedlund et al. unpublished) Bacillus sp. JF8 NR (Miyazawa et al. unpublished) Bacterium sp. NK3, NK2, NJ2 NR (Widada et al. unpublished) Pseudoalteromonas sp. EH-2-1 NR (Hedlund et al. unpublished) ndoAaAbAcAd Pseudomonas putida NCIB 9816 NR (Kurkela et al. 1988) Pseudomonas putida ATCC 17484 NR (Hamann unpublished) Pseudomonas fluorescens ATCC 17483 NR (Hamann unpublished) Pseudomonas sp. 30-2 NR (Panicker et al. unpublished) Thermoplasma acidophylum NR (Ruepp et al. 2000) Bordetella parapertussis 12822 NR (Parkhill et al. 2003b) Bordetella pertussis Tohoma I NR (Parkhill et al. 2003b) doxAaAbAcAd Pseudomonas sp. NR (Denome et al. 1993) pahAaAbAcAd Pseudomonas putida OUS82 chrom (Takizawa et al. 1994) Comamonas testosteroni H NR (Moser et al. 2001) Pseudomonas aeruginosa PaK1 chrom (Takizawa et al. unpublished) nagAaAbAcAd Ralstonia sp. U2 pWWU2 (Fuenmayor et al. 1998) bphAaAbAcAd Sphingomonas aromaticvorans F199 pNL1 C (Romine et al. 1999b) bpfAaAbAcAd Rhodococcus opacus NCIB12038 NR (Larkin et al. 1999) narAaAbAcAd Rhodococcus sp. 1BN NR (Andreoni et al. 2000) Rhodococcus sp. P200, P400 NR (Chen et al. unpublished) nidAaAbAcAd Mycobacterium vanbaalenii PYR-1 NR (Khan et al. 2001) phnAaAbAcAd Ralstonia sp. NI1 NR (Widada et al. 2002) Burkholderia phenazinium Hg8, Hg10, Hg16, Hg14 NR (Wilson et al. 2003) Burkholderia glathei Hg2, Hg4, Hg11 NR (Wilson et al. 2003) Cycloclasticus sp. A5 NR (Kasai et al. 2003) nsaAaAbAcAd Sphingomonas sp. BN6 NR (Conradt et al. unpublished) dntAaAbAcAd Burkholderia sp. DNT NR (Leungsakul et al. unpublished) Fluorene dioxygenase flnAaAbAcAd Terrabacter sp. DBF63 NR (Habe et al. unpublished) Phenanthrene dioxygenase phnAaAbAcAd Burkholderia sp. RP007 NR (Laurie et al. 1999a) Sphingomonas chungbukensis. DJ77 NR (Kim et al. unpublished) Alcalingenes feacalis AFH2 NR (Kiyohara et al. unpublished) adhAaAbAcAd Sphingomonas sp. P2 NR (Pinyakong et al. 2003b) Sphingomonas sp. AJ1 NR (Iwabuchi et al. unpublished) phdAaAbAcAd Nocardioides sp. KP7 NR (Iwabuchi et al. 1998)

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nidAaAbAcAd Mycobacterium vanbaalenii PYR-1 NR (Krivobok et al. 2003) Pyrene dioxygenase nidAaAbAcAd Mycobacterium vanbaalenii PYR-1 NR (Krivobok et al. 2003) pdoAaAbAcAd Mycobacterium sp. 6PY1 NR (Krivobok et al. 2003) Mycobacterium sp. S65 NR (Sho et al. unpublished) ‡ The dioxygenase gene operon: geneAa encodes the reductase, geneAb encodes the ferredoxin, geneAc encodes the large α- subunit of ISP, geneAd encodes the small β-subunit of ISP. * Location of the catabolic genes in the genome: chrom = on the chromosome, p = on a plasmid, NR = not reported; C = via conjugation transferable plasmid

Based on sequence similarity, the PAH-dioxygenases isolated from Gram negative Proteobacteria can be divided into three types which differ from taxa to taxa. Proteobacteria from the α-subcluster seem to have similar initial PAH-dioxygenases that differ from dioxygenases produced by members of the β-subcluster or the γ- subcluster. The initial PAH-dioxygenase genes of PAH-degrading α-Proteobacteria were found to be highly conserved among many Sphingomonas species such as S. yanoikuyae strains B1 and Q1, S. aromaticivorans F199, S. ‘agestris’ HV3, S. chungbukensis DJ77, S. xenophaga BN6, Sphingomonas sp. TNE12, Sphingomonas sp. P2 and Sphingomonas sp. EPA505 but divers for the 2 other Proteobacteria gene systems (Bastiaens 1998; Laurie et al. 1999b; Pinyakong et al. 2003a). Similarly, the phenanthrene dioxygenase catabolic genes sequenced from β-Proteobacteria Commamonas strains and Burkholderia strains capable of growing at expense of naphthalene and phenanthrene, showed high homology among each other (Goyal et al. 1996; Laurie et al. 1999a; Laurie et al. 1999b) but low homology with the naphthalene degradation genes conserved within the γ-Proteobacteria. Naphthalene dioxygenase genes are highly conserved among γ-Proteobacterium Pseudomonas strains (87-93% similarity) (Meyer et al. 1999) but clearly different from naphthalene dioxygenase genes purified from Sphingomonas, Mycobacterium or Rhodococcus strains degrading higher molecular PAHs (Hamann et al. 1999; Larkin et al. 1999; Laurie et al. 1999b; Meyer et al. 1999; Treadway et al. 1999). In addition, phylogenetic analysis has indicated that all initial aromatic dioxygenases from Gram positive bacteria form a subfamily of enzymes distinct from the dioxygenases from Proteobacteria (Meyer et al. 1999; Saito et al. 2000; Khan et al. 2001). The large subunit of the initial phenanthrene dioxygenase Phd (PhdA) from Nocardioides was 57% identical to large subunit of Nid enzyme (NidA) sequenced from phenanthrene and pyrene degrading M. vanbaalenii PYR-1 (Khan et al. 2001; Khan et al. 2002; Brenza et al. 2003; Krivobok et al. 2003). All dioxygenases sequenced from

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Mycobacterium strains were found to be highly homologous (minimum 98% identity) (Khan et al. 2001; Khan et al. 2002; Brenza et al. 2003; Krivobok et al. 2003).

The strong conservation of the nahAc homologs among the Gram negative naphthalene degrading isolates suggest in situ horizontal gene transfer (Herrick et al. 1997). Similarly, the high similarity and the exclusive presence of the nid genes in PAH-degrading Mycobacterium strains and not in non-degrading related strains of the same species, suggest a common origin of PAH-dioxygenase genes from Mycobacterium strains and that Mycobacterium strains obtained the nid genes later in evolution, possibly by horizontal transfer (Brenza et al. 2003).

Catechol-hydroxylating dioxygenases

In a second step, a dehydrogenase will transform the product of the oxygenation reaction, i.e., the cis-dihydrodiol to a dihydroxylated aromatic intermediate, i.e., a catechol (Cerniglia 1992; Juhasz et al. 2000b). These catechols may then be processed through either an ortho-cleavage type pathway (ring cleavage between the 2 hydroxyl carrying carbon atoms) or meta-cleavage type pathway (ring opening next to the hydroxyl carrying carbon atoms) (Figure 1-2) (Juhasz et al. 2000b). The final metabolites produced through both pathways (succinate, fumarate, pyruvate, acetate and acetaldehyde) will be further metabolized through central metabolic pathways for the synthesis of new cellular components or the production of energy and will be mineralized to CO2 and H2O. The ortho-pathway is catalyzed by intra-diol dioxygenases or catechol-1,2-dioxygenase and the meta-pathway by extra-diol dioxygenases or catechol-2,3-dioxygenase (C23O).

The majority of bacteria growing with PAHs as sole sources of carbon and energy follow the meta-cleavage pathway using a catechol 2,3-dioxygenase (C230) (Cerniglia 1992; Grifoll 1992; Boldrin et al. 1993; Monna et al. 1993; Resnick et al. 1996; Casellas et al. 1997; Meyer et al. 1999; Wattiau et al. 2001; van Herwijnen et al. 2003b). However, enzymes of both the meta- and ortho-pathway of catechol degradation were shown to operate in the process of naphthalene degradation by Pseudomonas strains (Kulakova et al. 1989; Kosheleva et al. 1997). Moreover, P. putida strains degrading naphthalene via the ortho-pathway were found to be more

- 25 - Chapter 1 competitive than the P. putida strains degrading naphthalene via the meta-pathway in mixed chemostat cultures on naphthalene (Filonov et al. 1997). In Gram negative bacteria, anthracene is mostly metabolized via a meta-cleavage pathway analogous to that of naphthalene metabolism to yield salicylate and catechol (Cerniglia 1984; Cerniglia 1992; Sutherland et al. 1995). In Gram positive bacteria, on the other hand, anthracene is mostly mineralized via the ortho-cleavage pathway through o-phthalate and protocatechuate (Dean-Ross et al. 2001; van Herwijnen et al. 2003a).

FIGURE 1-2 THE ORTHO- AND META- DEGRADATION PATHWAYS FOR AROMATIC RING CLEAVAGE OF A CATECHOL

Catechol 2,3-dioxygenase gene sequences have been determined for PAH-degrading bacteria of the genera Pseudomonas, Sphingomonas (Yrjala et al. 1998; Meyer et al. 1999) and Burkholderia (Laurie et al. 1999b). Based on sequence similarity, the catechol-dioxygenases isolated from Gram negative Proteobacteria can be divided into three types which differ from taxa to taxa. Proteobacteria from the α-subcluster seem to have similar catechol-dioxygenases that differ from dioxygenases produced

- 26 - Literature Review by members of the β-subcluster or the γ-subcluster. Except the catabolic genes for the lower fluorene degradation pathway of Sphingomonas sp. LB126, i.e., protocatechuate catabolic genes, which seemed to be more closely related to the genes previously found in lignin-degrading Sphingomonas sp. SYK-6 than to the corresponding genes of other PAH-degrading Sphingomonas strains (Pinyakong et al. 2003a; van Herwijnen et al. 2003b). It has been speculated that the ability of Sphingomonas strains isolated from the subsurface (deeply-buried sediments in the Atlantic ocean) to degrade a wide array of aromatic compounds including PAHs represent an adaptation for the utilization of sedimentary lignite, the major source of organic carbon for heterotrophic organisms in that environment (Fredrickson et al. 1999).

Location, organization and regulation of the PAH-degradation genes

In γ-Proteobacteria Pseudomonas strains, the naphthalene degradation genes have mostly been located in polycistronic operons (Burlage et al. 1989; Menn et al. 1993). However, the gene order of the catabolic genes of β-Proteobacteria Burkholderia and Commamonas is significantly different from the classical naphthalene degradation nah-like systems of Pseudomonas strains (Goyal et al. 1996; Laurie et al. 1999a). The genes for PAH-degradation in α-Proteobacteria Sphingomonas were often complexly arranged, i.e., the genes necessary for one degradation pathway were scattered over several operons and gene clusters (Feng et al. 1997). The metabolic pathways for different monocyclic and polycyclic aromatic hydrocarbon degradation were often linked through a grouped organization and co-regulation of the catabolic genes involved (Kim et al. 1999; Romine et al. 1999a).

The catabolic genes for aerobic oxidation of PAHs such as naphthalene and fluorene are localized in the chromosome or on plasmids. In γ-Proteobacteria Pseudomonas strains, the naphthalene degradation genes are often located on plasmids such as pNAH7 (Schell 1986; Burlage et al. 1989; Selifonov et al. 1991; Menn et al. 1993), pNPL-1 (Kulakova et al. 1989; Selifonov et al. 1991; Kozlova et al. 1999), pBS2 (Kulakova et al. 1989; Selifonov et al. 1991; Kozlova et al. 1999), pBS216 (Kulakova et al. 1989), pBS217 (Kulakova et al. 1989), pBS3 (Selifonov et al. 1991), pBS4 (Selifonov et al. 1991), pDTG1 (Simon et al. 1993; Park et al. 2002), pCg1 (Park et

- 27 - Chapter 1 al. 2003), pND6-1 (Li et al. unpublished) and pLP6a (McFarlane et al. unpublished). It has been shown that the Pseudomonas PAH-catabolic systems located on plasmids are thermo-sensitive as the catabolic plasmids were eliminated at 41-42 °C. Some of these catabolic plasmids had even an inhibition effect on growth of Pseudomonas strains at an elevated temperature, as plasmid free mutants could grow much better on elevated temperature (Kochetkov et al. 1983). Plasmids containing PAH degradation genes have also been isolated from Sphingomonas strains such as S. aromaticivorans F199 (the 184kb conjugative pNL1 plasmid) (Romine et al. 1999b) and Sphingomonas sp. KS14 (a >500kb mega-plasmid). The presence of large plasmids, but with unknown function, has also been demonstrated in many other PAH- degrading Sphingomonas strains (Bastiaens 1998; Fredrickson et al. 1999; Romine et al. 1999a; Bastiaens et al. 2000). So far no plasmids carrying catabolic genes have been identified in Mycobacterium strains (Bastiaens 1998; Bastiaens et al. 2000; Brenza et al. 2003). The genes for PAH-catabolism from Mycobacterium strains are therefore thought to be localized on the chromosome.

Although PAH-catabolic genes from Gram-positive and Gram-negative bacteria are phylogenetically different, they have nevertheless many similarities both in sequence, location and gene organization, suggesting that they have a common although distant, evolutionary origin (Harayama et al. 1992; Mason et al. 1992; Pinyakong et al. 2003a).

Conclusions

Microbial degradation is considered to be the major route through which PAHs are removed from contaminated environments and therefore is considered as feasible remediation technology. Currently, in situ and ex situ bioremediation techniques are, however, still very inefficient for removal of PAHs from contaminated soil. As any other technology engineered biological remediation of PAH-contaminated sites can only become successful if the active system is known and relative controllable. This requires more information about the identity of the degrading soil microorganisms and the enzymatic mechanisms they use. PAH-degrading microbial communities have been detected in many different contaminated habitats. Ecological analysis of microbial communities in PAH-

- 28 - Literature Review contaminated environments has shown that many different bacterial genera and species can be involved in the degradation process. The environment conditions and the type and concentration of PAHs seem to select for specific genera and species independent of the geographical location of the samples. Mostly Proteobacteria from the genera Pseudomonas (γ), Burkholderia (β) and Sphingomonas (α) or Corynebacterineae from the genera Nocardia, Rhodococcus and Mycobacterium seem to be involved in PAH-biodegradation in soil. The first group of Gram negative bacteria seems to be degrading preferentially lower molecular PAHs such as naphthalene and phenanthrene, while the second group of Gram positive bacteria is more specialized in the degradation of higher molecular PAHs such as pyrene. These PAH-degrading bacteria seem to have adapted to their hydrophobic PAH-substrates by showing low nutrient requirements and by making use of bioavailability promoting systems such as production of biosurfactants, high substrate affinity, mouth-like uptake systems or close contact biofilm formation. Mycobacterium species seem to be specialized in using hydrophobic sorbed or organic dissolved PAHs while Pseudomonas and Sphingomonas strains prefer aqueous liquid systems. Several aerobic microbial PAH-degradation pathways have been identified. The catabolic genes from Gram-positive and Gram-negative bacteria coding the dioxygenase enzymes are phylogenetically different, and are conserved on the genus level. In Pseudomonas and Sphingomonas strains, PAH-catabolic genes are often localized on conjugative plasmids while in Mycobacterium species they seem to be chromosomal. The genes of different genera have nevertheless many similarities both in sequence, location and gene organization, suggesting that they have a common although distant, evolutionary origin. The information on bacterial PAH-metabolic processes so far will allow the development of new tools that will enable researchers to analyze PAHs contaminated sites efficiently and to determine more rapidly strategies for treating them. For example, new nucleic acid probes and biologically based sensors could be constructed. It also provide the capacity to modify organisms for improved degradative performance through new metabolic pathway construction - either to complete a degradative process or to broaden its specificity to accommodate previously untreatable molecules - and to manipulate the genetic regulatory elements to improve efficiency.

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MICROBIAL BIOREMEDIATION OF PAH-CONTAMINATED SOILS

Technologies available for remediation of PAH-contaminated soil

To clean-up PAH contaminated soil and sludge different soil remediation technologies are available. Some of these technologies allow in field treatment (in situ) while others demand excavation (ex situ) with (off site) or without (on site) transport. The choice of the most appropriate remediation technique for a given polluted site depends on (A) pollutant characteristics, such as (i) the type and concentration of the PAHs, (ii) the type and concentration of the co-contaminant, (iii) the pollution history, and (B) soil characteristics, such as (i) pollution of top soil or/and unsaturated zone or/and aquifer, (ii) soil permeability for air and water, (iii) the location of the soil, (iv) soil temperature, pH, natural chemical composition and presence of natural terminal electron acceptors, (v) soil nature microflora and (vi) stability of soil structures.

Physico-chemical soil remediation

The conventional techniques used for remediation have been the excavation of the contaminated soil and transport it to a landfill or containment of the contaminated areas by capping. However, the first approach moves the contamination elsewhere and may create significant risks during the excavation, handling and transport of hazardous material. Additionally, land fill sites are increasingly expensive for disposal of the material. The ‘cap and contain’ method is only an interim solution since the contamination remains on site, requiring long term monitoring and maintenance of the isolation barriers.

A better approach is to completely remove the pollutants from the contaminated soil matrix or if possible, to destroy the pollutants by transforming them in innocuous substances. Mainly ex situ and off site thermal, physical and chemical extraction and decomposition techniques have been used. In the ‘thermal desorption’ technique the polluted soil is heated to 600 °C and PAHs are partially degraded by pyrolyse and transported from the solid phase to the vapor phase (Sutherson 1997). The airflow removing the PAHs is treated in a controlled combustion reaction completely

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oxidizing the PAH compounds to CO2 and H2O. ‘Soil washing’ is a combination of physical and chemical extraction methods to separate PAHs from the soil, or at least separate highly contaminated soil fractions from less contaminated fractions (Sutherson 1997). Water and chemicals are intensively mixed with the polluted soil, improving desorption and solubilisation of the PAHs in the water phase. Sometimes active carbon particles are mixed to enhance extraction of PAHs by sorption on the carbon particles. Organic substances and small soil particles are removed based on their particle size and density by pumping the soil slurry through a series of hydrocyclons and flotation beds. The pollutant concentrations in the ‘washed soil’ fraction are reduced below legal limits and can be reused while the separated fraction, highly enriched in PAHs, is incinerated or deposited. Such ex situ physico-chemical techniques are very effective in reducing contaminant concentrations but have several disadvantages, such as their technological complexity, the high cost for small-scale application, and the lack of public acceptance. Soil washing will generate a highly pollutend final soil fraction wich still needs to be dumped or incinerated. Incineration, has been protested as it may increase the exposure for both the workers at the site and nearby residents. Incineration will also result in combustion of all organic material and sterilization of the soil. The treated soil will loose all biological life and a large part of its economical value and can only be used in the construction industry. In addition, the large scale size of many contaminated areas makes the physico-chemical approach highly costly and unfeasible.

Alternatively, for some sites in situ and on site extractive remediation techniques can be used. In porous soils, the light molecular weight volatile fraction of PAHs can be removed from soil by soil vapor extraction combined with air infiltration. Clean air is pumped in the soil above (‘airventing’) or below (‘airsparging’) the ground water table under pressure while soil air is extracted from the dry zone under vacuum. In addition, a soil water extraction (‘pump and treat’) approach can be used to remove the water-soluble but non-volatile PAH fraction from contaminated aquifers in porous sandy soils. Pump-and-treat can be combined with the infiltration of water containing detergents to leach out the pollutants (‘soil leaching’ method). When high concentrations of PAHs are trapped in a non-aqueous phase (NAPL) floating on the water table, selective extraction of NAPL under vacuum can be used (‘slurping’ method) combined with air and water extraction techniques applied on the capillary

- 31 - Chapter 1 zone. ‘Chemical oxidation’ has proven to be an efficient in situ treatment for remediation of PAH-contaminated soil and ground water (Barton et al. 2000; Siegrist 2000). However, oxidation with peroxide, ozone, permanganate or ultrasonic radiation will only partially degrade the PAH-compounds.

Biological soil remediation

Bioremediation has emerged the last decennia as an alternative technology for the clean up of hydrocarbon contaminated soils (Vidali 2001). Bioremediation is defined as the process in which organic contaminants are biologically degraded under controlled conditions to an innocuous state, or to levels below concentration limits established by regulatory authorities (Vidali 2001). Microorganisms naturally present in the soil such as bacteria and fungi are stimulated to degrade or transform pollutants to less toxic compounds. As such, bioremediation is a relatively low-cost and low- technology approach, which generally has a high public acceptance and often can be carried out on site (Wilson et al. 1993) (Table 1-3). The efficiency of bioremediation is however site- and pollutant- dependent (Table 1-3). The range of contaminants on which it is effective is limited, the time scales involved are relatively long, and the residual contaminant levels may not always be appropriate (Raffi et al. 1994) (Table 1-3). In addition, considerable expertise may be required to design and implement a successful bioremediation program, due to the need to thoroughly assess a site for its suitability and to optimize conditions to achieve a satisfactory result (Table 1-3).

TABLE 1-3 ADVANTAGES & DISADVANTAGES OF BIOREMEDIATION

Advantages Disadvantages • high public appreciation • extrapolation from bench & pilot-scale • relatively low cost studies to full-scale field operations is very • can often be carried out in situ or on site difficult • (minimal disturbance, lower risks for • relatively long treatment time required contaminant distribution due to transport & • very much dependent on environmental handling) factors and contaminants mixture and • useful for a wide variety of organic dispersion contaminants complete destruction of the • limited to those compounds that are contaminant biodegradable

• residues are usually harmless (CO2, H2O, • residues may be more persistent or toxic biomass) than the parent compound Several ex situ bioremediation approaches have been developed to solve the contamination problem more rapidly (Table 1-4). These techniques involve the

- 32 - Literature Review excavation of contaminated soil from ground and its transport to a ‘bioremediation facility’. Soil remediation ‘bioreactors’ allow full control and optimization of the degradation process (Vidali 2001) (Table 1-4). Depending on the reactor type the contaminated water, soil or sludge is treated as pure liquid (aqueous reactor), in slurry with 5 to 40 % dry weigth (slurry reactor) or as dry soil (dry soil reactor or DSR). Extensive mixing conditions allow maximum interaction between bacteria, soil particles, contaminants and nutrients stimulating biodegradation. Full control of degradation parameters such as aeration, temperature, nutrient addition and bacteria inoculation could severely reduce the remediation time from months to weeks or days (Cookson 1995) (Table 1-4). In general, the rate and extent of biodegradation are higher in a bioreactor system than in in situ or ex situ solid-phase system because the contained environment is more manageable and hence more controllable and predictable. The use of ex situ bioreactors on lab or pilot scale has been found relatively effective for remediation of soil containing a complex PAH mixture (Wilson et al. 1993; Bastiaens 1998). For real field applications, the running costs for bioreactors are generally higher than other ex situ and in situ treatments (Figure 1-4). ‘Landfarming’, ‘biobeds’ or ‘biopiles’ are ex situ techniques where the contaminated soil is spread out on an impermeable underground (Vidali 2001). Supplements to neutralize soil pH and/or to improve soil texture (sand, straw, compost or wood chips) and sometimes extra nutrients and/or bacteria are mixed with the soil. Tilling of the biobeds on a regular time scale improves oxygen and nutrient distribution. In addition, water is percolated and air can be pumped through the soil to improve biodegradation conditions. To reduce atmospheric pollution by volatile hydrocarbons, biobeds can be placed indoors or covered with plastic to capture and treat the air. When high concentrations of non-hazardous organic additives such as manure, straw or agricultural wastes are mixed with the soil, elevated temperature can be reached in the biobed due to degradation of organic material and the term ‘composting’ may be used instead of landfarming (Semple et al. 2001). In general, these techniques have relatively low monitoring and maintenance costs and relatively high clean-up liabilities compared to other ex situ treatments (Vidali 2001) (Table 1-4) (Figure 1-4). Landfarming and composting has been successfully applied on a large scale to different soil types contaminated with low to middle distillated hydrocarbon mixtures such as diesel fuel, jet fuel and gasoline (Tien et al. 1999; Semple et al. 2001).

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However, biodegradation of PAHs by landfarming has been much less successful (Wilson et al. 1993). Landfarming methods could reduce PAHs contamination levels within a reasonable period of time but only PAHs with three or fewer aromatic rings were degraded (Wilson et al. 1993; Semple et al. 2001).

In addition, in situ bioremediation techniques can be applied to the soil and groundwater (Table 1-4). These techniques are generally the most desirable bioremediation option to remediate sites in use and/or sites with low-level contamination of surface soil over a vast area due to lower cost and the minimal disturbance avoiding excavation and transport of contaminants. The currently most applied engineered in situ bioremediation technologies include ‘bioventing’ and ‘biosparging’ (Radway et al. 1996) (Table 1-4). Through extraction and infiltration of air and water in the soil, it is possible to increase the bioavailability of the PAHs and the oxygen supply for the aerobic PAH-degrading bacteria in the vadose respectively water-saturated zone (Vidali 2001). ‘Biological permeable reactive barriers’ or ‘bioscreens’ can be installed at the borders of the contaminated site perpendicularly to the groundwater flow to contain and to treat the PAH contaminated ground water in order to protect downstream areas. The PAH- biodegradation is locally stimulated by the addition of nutrients and oxygen in the bioscreen zone. After passing through the bioscreen the ground water will be free of contaminants. As this technique requires no active pumping the operational costs are low (Figure 1-4). A new in situ technique currently in development combines the use of electrokinetics to transport water, ions and bacteria through PAH polluted clay soils to stimulate in situ biodegradation (Ho et al. 1995). Distribution control and natural attenuation, i.e., biodegradation without human interference, could be an alternative in situ bioremediation strategy (Table 1-4). Such passive in situ bioremediation processes can be economically very attractive but requires regular monitoring and longer times to reach desired pollutants levels (Figure 1-4).

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TABLE 1-4 SUMMARY OF BIOREMEDIATION STRATEGIES

Application Technology Benefits Limitations Ex situ Slurry reactors Rapid degradation kinetics Soil requires excavation Bioreactors Optimized environmental parameters Relatively high cost capital Enhanced mass transfer Relatively high operating cost Effective use of inoculants and additives Ex situ Landfarming Cost efficient Soil requires excavation Composting Low cost Space requirements Can be done on site Extended treatment time Need to control abiotic loss In situ Bioventing Most cost efficient Environmental constraints Biosparging Noninvasive Geological constraints Natural attenuation Relatively passive Extended treatment time Treats soil & water Monitoring difficulties

COST Slurry reactor Bioreactor Intensive landfarming Extensive landfarming

Active In situ bioremediation Passive In situ bioremediation

TIME

FIGURE 1-4 RELATIONSHIP BETWEEN TREATMENT TIME AND COST FOR BIOREMEDIATION METHODS

Assessment and monitoring of biodegradation potential

A first step in a bioremediation strategy would be ‘a feasibility assessment and field evaluation’. The microbial community naturally present in the polluted soil is analyzed and tested for its potential to degrade the PAHs under predetermined laboratory conditions to identify limiting factors and recommend ways to mitigate these limitations in the field (Balba et al. 1998).

The intrinsic metabolic potential of a contaminated soil is mostly assessed by degradation tests in the lab using laboratory microcosm systems (Heitzer 1993). Contaminated soil samples are incubated in the lab under different conditions to

- 35 - Chapter 1 screen bioremediation treatments and select the most appropriate strategy for large scale application (Balba et al. 1998). These microcosms can vary in complexity from simple static soil jars to highly sophisticated mini bioreactor systems containing dry soil or soil slurry. There are different ways to assess the PAH-degradation potential in such microcosm systems.

A first method is by measuring the total dehydrogenase activity in the soil as biological oxidation of organic compounds is generally a dehydrogenation process that is catalyzed by dehydrogenase enzymes (Balba et al. 1998). The most widely used method for the determination of soil dehydrogenase activities is the colorimetric method, involving the used of 2,3,5-triphenyl tetrazolium chloride (TTC) which acts as an electron acceptor for many dehydrogenase enzymes and which is reduced to form the red compound triphenyl formazan (TPH). The intensity of the red color produced from the dehydrogenase assay is a good index for microbial activities within the tested soil. However, several soil compounds such as nitrate, nitrite and ferric ions may interfere with the dehydrogenase activity by ions acting as alternative electron acceptors.

An alternative method to assess the total biological activity in the sample is by measuring the total O2 consumption or total CO2 production rate (respirometry). This approach provides rapid, relatively unequivocal time-course data suitable for testing different biological treatment options, such as the effect of nutrient supplementation, microbial inoculation, etc. on microbial activity. To investigate more specifically the PAH-degradation potential, the mineralization of freshly added 14C-labeled PAHs is measured (radiorespirometry) (Spain et al. 1980; Spain et al. 1983; Grosser et al. 1991; Carmichael et al. 1997; Reid et al. 2001). To assess the relative contribution of bacteria and fungi to the mineralization of PAHs, selective inhibitors such as cycloheximide (fungal inhibitor) or a mixture of penicillin B and tetracycline (bacterial inhibitors) can be added to identical sets of respirometers, along with the 14C-labeled PAHs (Macleod et al. 2002). However, added inhibitors are only limited effective due to natural resistance to them and their short period of action due to deactivation and sorption to the soil matrix (active for 2-4 days depending on the soil type) (Anderson et al. 1973; Anderson et al. 1975; Stamatiadis et al. 1990). In addition, specific removal of competitors by adding selective inhibitors may result in

- 36 - Literature Review a faster growth of surviving microorganisms (Anderson et al. 1975). In addition, also the residual concentrations of PAHs and their degradation products can be monitored during treatment in a sub-sampling or a batch set-up (liquid or gas chromatography).

A more direct approach to asses the PAH-degradation potential of a soil, is to look for the presence of bacteria with known PAH-degrading capacities or known bacterial PAH-degradation enzyme systems. Culture-dependent detection of bacteria is mostly done by microbial enumeration by plate counting on appropriate media, for example media with PAH as sole C-source (Balba et al. 1998). However, the agar plate microbial-counts technique has several limitations particularly when dealing with non-culturable microorganisms in soil. Culture-independent molecular methods based on the detection of bacterial DNA, RNA or enzymes involved in PAH-degradation are often preferred to give more rapidly information about the presence and activity of PAH-degrading in the soil. DNA or RNA extracted from soil is used as template in specific PCR reactions. Specific 16S rDNA based detection methods allow to screen the soil for the presence of unknown types of bacteria in general or certain groups of bacteria in specific. The catabolic DNA coding initial PAH ring-hydroxylating dioxygenases and the catechol- cleaving dioxygenases have been used as targets to detect the presence of PAH degradation potential at the DNA level. A limited number of specific or degenerate probes and PCR primers have been designed based on known DNA sequences encoding the initial ring-hydroxylating dioxygenases and the C23O meta ring- cleaving dioxygenases of certain species (Hamann et al. 1999; Meyer et al. 1999). However, PAH degradation enzymes can be quite different at the DNA level without affecting their function. Many catabolic genes with different sequence may exist and may not be detected (Widada et al. 2002), leading to an underestimation of the degradation potential. It is therefore necessary to gain more sequence information for different PAH-degrading strains of different taxa to establish a universal or group specific PCR and RT-PCR protocols for the detection of PAH degradation genes and mRNA, respectively (Meyer et al. 1999; Widada et al. 2002). The total size and activity of a bacterial community can be assessed by measuring the 16S rRNA:rDNA ratio of the cells as they grow in the soil (Ka et al. 2001). To follow the activity of certain species or strains, this technique can be combined with specific 16S rDNA oligonucleotide hybridization probes.

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Limitations and strategies to improve engineered bioremediation of PAH- contaminated soil

Currently, mostly the soil is basically treated as a ‘black box’ during bioremediation. Not much is known about the key organisms needed for PAH-biodegradation and the specific needs of PAH-degrading bacteria with respect to nutrition and environmental conditions for their optimal activity in the soil environment. Bioremediation can only be effective where environmental conditions permit microbial growth and activity. Bioremediation therefore often involves the manipulation of environmental parameters to allow microbial growth and degradation to proceed at a faster rate.

Bioaugmentation

The first criterion for successful bioremediation is the presence of microorganisms in the soil that actually are capable of degrading the pollutant, i.e., microorganisms that posses appropriate enzymatic systems to catalyze the degradation reactions. It is assumed that whenever the pollutants are biodegradable and the soil conditions are favorable, suited bacterial communities to do the job will most likely have naturally evolved (Ide et al. 1996). Most characterized PAH degrading isolates are able to degrade 1 or a few different PAHs. However, consortia of different bacteria or maybe new engineered strains with as wide spectrum of degradation enzymes are needed for the degradation of complex PAH mixtures in soil (Bouchez et al. 1995; Bastiaens 1998). A cell density of 106 to 108 CFU g-1 dry weight is mentioned as a minimum cell concentration to obtain suitable biodegradation rates (Ramadan 1990). If not ‘sufficient’ bacteria capable of mineralizing PAHs are present in the soil, a bioaugmentation strategy can be an option, i.e., the addition of pollutant degrading bacteria indigenous or exogenous to the contaminated site.

The positive effects of bioaugmentation on small scale are reported many times (Grosser et al. 1991; Hickey et al. 1993; Weir et al. 1995; el Fantroussi et al. 1997). Different authors have shown in laboratory and some pilot scale experiments the successful biodegradation of artificially PAH-contaminated soils by the addition of PAH degraders belonging to the genera Mycobacterium and Sphingomonas. 2- methylnaftalene, phenanthrene, pyrene and benzo(a)pyrene degradation was increased

- 38 - Literature Review by the inoculation of sediment and water microcosms with M. vanbaalenii PYR1 (Heitkamp et al. 1989). The reintroduction of 107 to 108 Mycobacterium sp. PAH135 cells per gram of soil could significantly increase the mineralization of pyrene (Grosser et al. 1991). The degradation of pyrene in soil was enhanced 10 times by adding a Mycobacterium gordonae-like strain or a Sphingomonas paucimobilis strain (Kästner 1998). A fluorene degrading Sphingomonas strain was succesfull inoculated in a ‘Dry Soil Reactor (DSR)’ containing 70 kg of a non sterile PAH-contaminated soil (70 % dry weight) and allowed relatively fast PAHs removal in a relatively short treatment time (Bastiaens 1998). The high degradation capacity and the bioavailability promoting characteristics make consortia of Sphingomonas and Mycobacterium strains especially suited for bioaugmentation of PAH polluted soil.

Reports of successful in situ or ex situ large scale bioaugmentation however, are less numerous. Points limiting the use of bioaugmentation in real site bioremediation are (i) the inefficient large scale cultivation of the bacteria to inoculate, (ii) the inefficient inoculation and distribution of the bacteria in the soil, and (iii) the limited colonization, survival and activity of the inoculated bacteria under the given soil or remediation conditions. To treat large amounts of polluted soils huge amounts of bacteria are needed. Therefore inoculum bacteria need to be efficiently and economically cultivated in large volume batch bioreactors to obtain sufficient biomass. This asks for cheap and well defined media satisfying the needs of the selected inoculum bacteria. However, using rich media allowing quick growth to high densities migth create bacterial cells that have difficulties to adapt and survive the harsh oligotrophic soil environment. Cultures pregrown on or in the presence of the target pollutants containing actively working enzyme machinery would be far more suited. Very little research has been done to develop good propagation conditions for large scale inoculation of bacteria in soil. So far, almost exclusively batch cultures have been used, but continuous cultivation could be economically more interesting. In addition, in most bioaugmentation cases only 1 single pure strain is used instead of a well adapted consortium of several strains attacking mixed pollutions. Using mixed inoculum cultures increases the need for efficient propagation methodologies. Bioaugmentation attempts were often also lacking a good inoculation methodology. Lab scale and pilot scale experiments have shown that the introduction of bacteria in

- 39 - Chapter 1 real environment can lead to rapid reduction of viability and activity (Roszak et al. 1987; Ramadan 1990; Weissenfels et al. 1992; Weir et al. 1995; Kästner 1998). Many inoculation parameters can influence the behavior of the added bacteria such as (i) the size of the inoculum, (ii) the inoculation medium and procedure, and (iii) the inoculum distribution in the soil. It was found that small inocula were incompetent (Ramadan 1990; Weir et al. 1995). In many lab experiments degradation activity increased linear with the inoculum size (Goldstein et al. 1985; Roszak et al. 1987; Kästner 1998). In general, cell concentration of 107 to 108 cells per gram soil is presented as the inoculum size limit for positive results in bioaugmentation. Bacteria can be added to the soil in liquid medium through simple sprinkling or mixing or could be added immobilized on for example polyurethane foam (Manohar et al. 2001). Mostly inoculum cells are suspended in buffers or media but not without any risks. It has been shown that the introduction of PAH-degrading bacteria suspended in a mineral medium had a negative influence on the degradation capacity of the indigenous and introduced microflora, while inoculation as a pure aqueous suspension did not affect the degradation activity (Kästner 1998). Immobilization has been reported to enhance the survival without decreasing the activity of added bacteria in soil and waste water during bioremediation processes. Sphingomonas cells bound to alginate or chitosan beads were successfully re-used several times in a lab- scale bioreactor for the treatment of nonylphenol contaminated industrial wastewater (Fujii et al. 2000). In addition a good mixing is critical for the uniform distribution of the cells and for the efficiency of the remediation process. For in situ bioaugmentation, subsurface irrigation techniques have proven to be much more efficient than surface irrigation as a microbial delivery tool (Mehmannavaz et al. 2002). Studies have shown that under natural conditions most motile endogenous and added soil bacteria have a limited mobility. For example several 2,4-dichlorphenol or p-nitrophenol degrading motile Pseudomonas strains were incapable to move through the pores of the soil matrix to locations with higher polluent concentrations (Goldstein et al. 1985). As a consequence, pollutants will only be degraded in the near vicinity of the inoculation points.

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After introduction, the inoculum bacteria will have to survive in the soil environment. The main cause for quick reduction in cell number in non sterile soil is probably the lack of competitiveness of many pure strain inocula adapted to lab cultivation conditions (Goldstein et al. 1985; Vidali 2001). Inocula will have to compete with the indigenous microflora for nutrients or colonization of niches. Only bacteria with high substrate affinity (specific biodegrading enzymes, biosurfactants, etc.) will be able to grow on poorly bioavailable pollutants such as PAHs in soil environments. It has been shown that the success of bacterial inocula to degrade PAH in soil was also depending on the concentration of organic material limiting PAH-bioavailability (Weissenfels et al. 1992). For some inoculum bacteria, polluent concentrations may be too low to deliver enough carbon and energy to sustain growth or to induce the enzymatic machinery needed for polluent mineralization (Goldstein et al. 1985; Ramadan 1990; Colores et al. 1999). On the other hand, high pollutant concentrations could inhibit biodegradation through nutrient- and oxygen depletion or through toxic effects of the pollutant on the inoculum (Leahy et al. 1990; Volkering et al. 1995).

Biostimulation

Apropriate PAH-degrading microorganisms in contaminated soil are not necessarily present in numbers sufficient for bioremediation of the site within a reasonable time period. The slow rate of PAH-biodegradation in soil in comparison to degradation rates in laboratory culture conditions, suggests that microbial activity is seriously constrained in the natural environment. The inherent properties of the PAH compounds and the soil properties are seriously affecting the biodegradation (Manilal et al. 1991; Wilson et al. 1993) (Table 1-5). To solve this problem and improve bioremediation efficiency, a better understanding of the interaction between the soil environment, the PAH-compounds and the biocatalysts is needed. For successful bioremediation of PAH-contaminated sites a system must be created in which the right soil microorganisms can be promoted to create and maintain sufficient biomass and to be metabolically active. In addition, process control is not only necessary to produce sufficient bacterial biomass but also to direct them towards optimal PAH- biodegradation activity.

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Nutrient availability for PAH-degrading bacteria in soil

In the environment, bacteria depend on the availability of several elements or nutrients in order to allow the cells to survive and build up new biomass (Baker et al. 1994; Sutherson 1997). There are three categories of nutrients based on the quantity and essential need by the microorganism, i.e., macro-, micro-, and trace nutrients. The macronutrients carbon (C), nitrogen (N) and phosphorus (P) are known to comprise 50%, 14% and 3% dry weight, respectively, of a typical microbial cell (Vidali 2001). The micronutrients sulfur (S), potassium (K) and sodium (Na) comprise 1% dry weight and calcium (Ca), magnesium (Mg) and chloride (Cl) 0.5% dry weight of a cell but play important roles in membrane and enzyme stability (Bailey et al. 1986). The most common trace elements are iron (Fe), manganese (Mn), cobalt (Co), copper (Cu) and zinc (Zn). These trace nutrients are only present in low quantities in the cells but are essential for enzyme functioning.

Based on this approach, it was suggested that optimal nutrient mixes for biomass formation should have an overall C/N/P-ratio of approximately 120/19.4/3.9 [w/w/w] (Paul et al. 1989). However, in real field situations mostly the C/N/P-conditions in soil are not optimal for the growth and activity of bacteria. Organic and inorganic substrates (carbon, nitrogen, phosphate, sulfur) can be added ex situ or in situ to balance the Carbon/Nitrogen/Phosphorus concentration ratio in soil. A C/N/P-ratio of 120/10/1 [mg], has been reported as optimal for cell growth on contaminants in general in a review about bioremediation of soil contaminated with PAHs (Wilson et al. 1993) (Table 1-5). More specific for optimal PAH-degradation in soil, an optimal overall C/N/P ratio of 120/2/0.15 [mg] was mentioned. Lately, most researchers recommend for bioremediation applications an optimal C/N/P mole-ratio of 100/10/1 [mole], i.e., a 120/14/3 ratio [mg] (Hoeppel et al. 1994; Bouchez et al. 1995; Cookson 1995).

• PAHs as carbon source

The principle of PAH-biodegradation is that heterothrophic soil microorganisms will use the carbon supplied by the PAHs for growth and energy. As exception to what

- 42 - Literature Review was previously said, carbon is the only substrate that is mostly needed at higher quantities then the chemical composition of the cell would predict. In general, a simple growth substrate can sustain a microbial yield of 60%, meaning that from 100mg of C only 60mg is incorporated into biomass and the other 40mg is used for energy production and transformed into CO2 (Paul et al. 1989). However, C from most pollutant compounds is mostly not substantially incorporated into microbial biomass, i.e., a larger fraction is mineralized to CO2 for the production of energy (Paul et al. 1989). This leads to higher pollutant concentrations needed to sustain microbial biomass (Paul et al. 1989).

The limited bioavailability of the PAH-compounds itself is believed to be the major bottleneck for efficient biological treatment of PAH-contaminated soil (Weissenfels et al. 1992; Erickson et al. 1993; Luthy et al. 1994; Beck et al. 1995; Würdemann et al. 1995). The rate at which microorganisms can mineralize hydrocarbons depends on (i) the rate of transfer of the compound to the microorganism and (ii) the rate of uptake and metabolism of the compound in the cell (Bosman et al. 1997) (Figure 1-5). Currently, most researchers believe that microorganisms can use PAHs only in dissolved state. Consequently, PAH solubilisation (desorption) and/or diffusive transport through the aqueous solution to the cell surface will affect PAH- biodegradation (Figure 1-5). For naphthalene and phenanthrene it was indeed shown that the growth rates of PAH-degrading bacteria were independent of the total PAH surface area presented to the bacteria, i.e., independent of the amount of solid-PAH, but limited by the solubility of the PAH compounds (Bouchez et al. 1995). It was shown that limited solubilisation and transport caused very slow growth and linear growth curves, indicating mass-transfer limited conditions, for bacteria that grew on 3-chlorodibenzofuran (Harms 1995). Based on this dissolved-based utilization, bioavailability and biodegradation of PAHs in soil is seriously limited by their hydrophobicity and concurrent sorption effects. PAHs tend to ad/absorb to the soil organic matter or accumulate strongly in non aqueous phase liquids. Sorbed PAHs desorb slowly and are therefore believed to be less available for biological uptake and degradation (Volkering 1992; Weissenfels et al. 1992; Volkering et al. 1993). The bioavailability of hydrocarbons in soils decreases with increasing molecular mass of the component (Cerniglia 1992) and with increasing soil hydrocarbon contact time, i.e., aging (Bauer et al. 1985; Weissenfels et al. 1992; Hatzinger et al. 1995; Macleod

- 43 - Chapter 1 et al. 2000; Reid et al. 2000). As such, even massive contaminations may be oligothrophic from a microbe’s perspective. Theoretical considerations show that for extremely hydrophobic contaminants as PAHs, the low solved available pollutant concentrations may not be sufficient to sustain the population size or to induce the degradation enzymes needed to achieve environmental clean-up (Goldstein et al. 1985). However, not all the researchers completely agree with this hypothesis and suggest that sorbed compounds are available to microorganisms without prior desorption (Guerin 1992; Volkering 1992; Mihelcic 1993). Some researchers state that the sorption of PAHs as thin films to the soil particles can lead to a better degradation by bacteria in comparison to the crystalline PAHs, due to increased substrate surface. The positive interaction between increasing substrate surface and a higher degradation and growth rate, due to higher mass transfer rates, has been described many times for mixed and pure cultures (Thiem et al. 1994; Tongpim et al. 1996; Bastiaens 1998).

FIGURE 1-5 MICROBIAL UPTAKE OF A SORBED POLLUTANT

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It is known that many PAH-degrading bacteria use original strategies to enhance bioavailability of the highly hydrophobic compounds. Some bacteria can increase the transport rate to and the uptake rate in the cells so that biodegradation rates exceeds the normal dissolution rates of the PAHs from solid or organic-solved state (Manilal et al. 1991; Bouchez et al. 1997; Bastiaens 1998; Tang et al. 1998; Garcia-Junco et al. 2001a; Wick et al. 2001; Carcia-Junco et al. 2003). These bioavailability promoting properties are (i) the excretion of biosurfactants to solubilize the PAH (Deziel et al.

1996; Willumsen et al. 1997; Yuste et al. 2000; Garcia-Junco et al. 2001a), (ii) high specific substrate affinity to increase PAH uptake rate (Button 1985; Wick et al. 2002a), (iii) attachment to the hydrophobic PAH crystal, the sorbed PAH, or the degradable or non-degradable organic phase solvent containing the PAH to decrease the cell–molecule distance (Ortega-Calvo et al. 1994; Harms 1995; Tongpim et al. 1996; Bouchez et al. 1997; Garcia-Junco et al. 2001a; Wick et al. 2002a; Wick et al. 2002b; Rodrigues et al. 2003). Up till now, still little is known about the physiological and biochemical mechanism involved in bacterial uptake of pollutants in the sorbed state. Even the specific physico-chemical characteristics of the substrate itself seem to determine the bacterial strategy regarding uptake, as some PAH induces biofilm formation and other not in the same strain capable of growing on both (Rodrigues et al. 2003). For some PAH-degrading Mycobacterium strains, there are strong indications based on electron microscopic observations that they posses special mechanisms to make very close contact with PAH-crystals (Wick et al. 2001; Wick et al. 2002a). As such, the persistence/bioavailability of PAHs in the environment thus not only depends on the physical and chemical characteristics of the PAHs, the composition and chemical characteristics of the sorbent but also on the specific properties of the PAH-degrading microorganisms present.

During engineered bioremediation the limitations of PAH-solubility can be reduced or solved in different ways. A chemical pretreatment consisting of an oxidation of the PAH structure with permanganate or ultrasonic radiation can partially degrade the PAH-molecules and enable microorganisms to metabolize them at a rate suitable for remediation purposes (Barton et al. 2000). The addition of synthetic nonionic surfactants, such as Triton X-100, Tergipol NPX, Brij 35 Igepal CA-720 or the oleophilic fertilizer Inipol EAP-22, in concentrations above their CMC to soil could increase the apparent solubility and therefore the

- 45 - Chapter 1 bioavailability and the biodegradation of PAH-compounds. However, experimental observations on the effects of surfactant addition on microbial degradation of hydrophobic hydrocarbons are not consistent (Aronstein et al. 1991; Edwards et al. 1991; Laha et al. 1992; Wilson et al. 1993; Efroymson et al. 1995; Volkering et al. 1998). The application of a surfactant has been reported to be beneficial (Volkering et al. 1995; Grimberg et al. 1996) or detrimental (Thibault et al. 1996) to microbial substrate utilization rates and growth yields. Possible beneficial effects are (i) the increase of aqueous solubility of the hydrocarbons, (ii) the reduction of interfacial tension that promotes formation of more interfacial area, (iii) the emulsifying action of the surfactant that overcomes interfacial area limitation and that permits effective contact between cells and substrate, or (iv) the liposome facilitated substrate transport through the microbial cell wall. Possible causes of negative effects could be (i) the toxicity of the surfactants to the bacteria, (ii) the sorption of the surfactant onto the soil, (iii) the CMC effect, i.e., the unavailability of the PAHs trapped in the micelles to the bacteria, (iv) the bacterium-surfactant interactions that effect cell membranes or prevent the bacteria to adhere to the hydrophobic hydrocarbon surfaces, or (v) the preferential metabolisation of nontoxic organic surfactants. In addition, provision of high surfactant concentrations on large scale is relative expensive and therefore preferably avoided. An alternative way to meet the limited C-source availability in PAH-contaminated soils could be the addition of extra C-substrates to the soil to maintain sufficient biomass and an active soil population. Historically, Broadbent and Norman (Broadbent et al. 1946) reported the stimulation of mineralization of one compound by added extraneous C-substrates already in 1946 and coined the term ‘priming effect’ to describe it. Non-inducer organic nutrients added to the soil could stimulate the soil microbial community and support co-metabolic transformation of PAHs. Shen and Bartha (Shen et al. 1996) demonstrated that the addition of glucose during bioremediation tests can stimulate the mineralization of organic carbon in soil. The stimulation of survival and PAH-degradation in soil by addition of extra C-substrates like skim milk has been reported by Weir (Weir et al. 1995) for a seeded Pseudomonas sp. that mineralizes phenanthrene. However, most researchers found that easy metabolisable organic nutrient amendments such as glucose or amino acids had no effect or interfered negatively with the degradation activity (Goldstein et al. 1985; Swindoll et al. 1988; Heitkamp et al. 1989; Carmichael et al. 1997). The supplements

- 46 - Literature Review probably limit the metabolisation of the PAHs because of rapid growth of non-PAH degrading microorganisms which out compete the PAH-degrading bacteria. Alternatively, the inhibitory effect could be due to ‘diauxic growth’, i.e., the preferential use of the supplement as carbon source above the pollutant by the same microorganisms. However, diauxic growth could be prevented if the substrates are added in multiple small doses (Harder et al. 1982). At low substrate concentrations (oligotrophic conditions) mixed-substrate growth is more common then diauxic growth (Kovarova et al. 1998). To stimulate a certain group of microorganisms, sometimes some specific organic pathway intermediates, i.e., inducers, are added. The survival of six naphthalene- degrading strains was significantly increased by adding salicylate, an intermediate in the naphthalene degradation pathway, to non-sterile soil samples (Ogunseitan et al. 1991). The biodegradation rate of naphthalene and phenanthrene in artificially contaminated soils was enhanced by addition of salicylate, while the degradation of pyrene was enhanced by the addition of gentisate, phtalate, cinnamate and propionate (Carmichael et al. 1997). These supplements had no effect however on PAH- degradation in natural historically contaminated soils or soils containing high PAH- concentrations. Moreover, at high supplement concentrations inhibition of degradation was observed due to accumulation of upstream metabolites. Steffensen and Alexander (Steffensen et al. 1995) suggest that the effect of one biodegradable substrate on the metabolisation of the second substrate is mostly related to the competition for inorganic nutrients in a mixed bacterial community.

• Inorganic macronutrients: Nitrogen and Phosphorus sources

Microorganisms need besides organic compounds as carbon sources (C) also macronutrients such as nitrogen (N) and phosphorus (P) for cell growth. Competition for nitrogen by plant roots, mycorrhizal fungi and microorganisms keep inorganic N and P concentrations in soil low. It is even suggested that the competition for inorganic nutrients among the microbial population can reduce the number of active PAH-degrading bacteria to an insufficient level, since the slower growing pollutant degrading bacteria are outcompeted (Cerniglia 1992; Steffensen et al. 1995). Nitrogen + - (N) is primarily used for cellular growth (NH4 or NO3 ) and as an alternative electron

- 47 - Chapter 1

- acceptor (NO3 ) instead of oxygen. Bacteria will use nitrogen to build amino acids and proteins for catabolism and to incorporate it in the peptidoglycan of their cell walls. In most soils, ammonia released during soil organic matter decomposition is rapidly transformed to nitrate through microbial nitrification processes. Phosphorus (P) is needed in the bacterial cell for the production of nucleic acids, phosphate sugars and phospholipids in the cell membrane. Phosphorus exists in nature in a variety of organic and inorganic forms but primarily in either insoluble or poorly soluble inorganic forms. The P-solubility in soil is regulated by a complicated equilibrium controlled by ion-ion association reactions, pH-effects and soil texture. Phosphorus tends to absorb to clay minerals. It exists mainly as apatites, with the basic formula

M10(PO4)6X2. Commonly the mineral (M) is calcium, iron or aluminum and the anion (X) is a fluor, cloor, hydroxy or carbonate ion (Paul et al. 1989). These bound forms are poorly extractable and therefore also considered as poorly bioaccessible. The portion that is extractable with water, diluted acids or bicarbonate solutions is designated as ‘available’ for uptake by living organisms. The concentration of free phosphorus in the soil solution is mostly of the order 0.1-1 ppm.

Addition of inorganic nutrients to balance N- and P-concentrations with the CPAH- concentrations in soil has been practiced to eliminate possible nutrient limitations in soil. Nitrogen is the nutrient most commonly added during bioremediation. Mostly nitrogen is added as urea (NH2CONH2), also called carbamide or carbonyldiamide, or as ammonium chloride (NH4Cl), but is also supplied as an ammonia-/nitrate- salt (e.g.

NaN03) or ammonium nitrate (NH4NO3). All these forms are readily assimilated in bacterial metabolism. However, many studies have indicated that ammonium, already being in the reduced state required for incorporation into amino acids, is preferred to the oxidized form nitrate which first has to undergo reduction. Even low levels of ammonia often repress the enzymes required for nitrate reduction (Paul et al. 1989). Nevertheless, some features of the supplements may be important depending on the soil situation. The application of urea in the unabsorbed form such as in manure will lead to high volatilization. Surface manure in the fields may lose up to 50% of its + nitrogen due to volatilization (Paul et al. 1989). The ammonium ion (NH4 ) creates an increased oxygen demand due to nitrification processes. In addition, while the anionic nitrate is very soluble and freely mobile in soil solution, the cationic ammonium is retained by soil cation exchange sites on clays (Bohn 1985). Ammonia also reacts

- 48 - Literature Review

with soil organic matter to form quinone-NH2-complexes. In addition, plants prefer ammonia to nitrate and thus ammonium can also be lost to plants. Nitrate is more susceptible to losses through leaching and denitrification (Paul et al. 1989). Fertilizer + nitrogen, added as urea (NH2CONH2), ammonia (NH3) or the ammonium ion (NH4 ) form, is also subject to nitrification (Paul et al. 1989). Gaseous nitrogen oxide (N2O) addition under high vapor pressure has been used as an alternative to distribute nutrients better in situ throughout soil (Radway et al. 1996; Bogan 2001). This new approach has been utilized in a patent process but it has not, however, been documented as a means of enhancing the remediation of PAH-contaminated soils (Bogan 2001).

Phosphorus is the second most commonly added nutrient in bioremediation and is supplied to serve as a source for cellular growth. As additional inorganic phosphor- source mostly solid monophosphate salts (also called orthophosphate salts) (e.g.

KH2PO4, K2HPO4, K3PO4) or polyphosphate salts (also called pyrophosphate salts)

(e.g. KH3P207) with potassium or sodium are added. Gaseous phosphate under the form of triethylphosphate (C3H9O4P) (TEP) and tributylphosphate (C12H27O4P) (TBP) have been used as an alternative for liquid or solid phosphate amendments (Radway et al. 1996; Bogan 2001). Although mildly toxic and corrosive irritants, TEP and TBP are considered the safest phosphorus compounds, which can readily be gasified. When added to soil, phosphate quickly absorbs, however, to iron and aluminum oxide surfaces and may even form precipitates with iron, aluminum, manganese and calcium (Brady 1990). Such reactions will limit its transport and causes it to be unavailable for biological activity (Johnson et al. 1990; Aggerwal et al. 1991; Ward et al. 1999). The addition of potassium phosphate may also accelerate the cleavage of hydrogen peroxide (H2O2) sometimes added simultaneously to the soil as an oxygen source for aquifer bioremediation. If the H2O2 cleaves immediately, this may generate microbial colonies growing only near the injection well and the oxygen source may be depleted before it reaches the contaminated zone (Ward et al. 1999). In most geochemical environments, 10 mg l-1 is the maximum orthophosphate addition to avoid significant precipitation, peroxide cleavage or toxicity (Riser-Roberst 1998).

- 49 - Chapter 1

The use of inorganic N and P supplements have been tested with a variety of organic pollutants, pure and mixed microbial cultures and environmental matrices. Unfortunately, the effects of nutrients on pollutant degradation rates in soil are very inconsistent. Addition of inorganic nutrients have been found to shorten the adaptation period for microbial degradation (Wiggins et al. 1987; Swindoll et al. 1988) or to increase the extent and rate of microbial metabolism of some pollutants in soil and water (Bossert et al. 1984; Swindoll et al. 1988; Manilal et al. 1991; Alexander 1994; Baker et al. 1994). Sometimes stimulation of pollutant degradation rates only appeared until days or weeks after nutrient addition (Jobson et al. 1974; Bossert et al. 1984). Many articles report no apparent effect on metabolism (Swindoll et al. 1988; Johnson et al. 1990) or even a decreased metabolism in other cases (Johnson et al. 1990; Manilal et al. 1991; Morgan et al. 1992). Lag time, initial rate, and degradation extent were affected singly or as a group by nutrient additions (Thorton-Manning et al. 1987; Swindoll et al. 1988). Moreover, in several reports there is a significant difference between the effects caused by the addition of N-salts, P-salts or both (Swindoll et al. 1988; Manilal et al. 1991). Phosphorus might be more often the limiting macronutrient in soils. A vague trend could be that P-supplements have more often a positive effect while N-supplements cause more frequently inhibition of biodegradation of organic pollutants. Inorganic salts of nitrogen and phosphorus can be very effective in closed systems but have the tendency to wash out in real outdoor field applications (Leahy et al. 1990). Very few or non overriding generalizations emerge from the studies that could deliver some practical information for real field applications. Differences in the bioavailability of nutrients added to soil may explain a portion of the observed differences in the response of degradation rates to nitrogen and phosphorus supplements. Just as sorbed carbon substrates are generally considered to be unavailable to soil microbes, sorbed inorganic nutrients also may be less bioavailable than nutrients dissolved in the soil solutions. Morgan and Watkinson (Morgan et al. 1992) suggested that high concentrations of supplements could lead to inhibition of microorganisms that are used to live in oligothrophic soil environments. Another important conclusion from all these studies may be that the responses to nutrient addition are largely depending on the soil that is used.

- 50 - Literature Review

• Micronutrients & Trace-elements

Next to macronutrient-concentrations, also micronutrients like sulphur (S), calcium (Ca), and magnesium (Mg) and trace-elements like the metals iron (Fe), manganese (Mn), cobalt (Co), copper (Pb) and zinc (Zn) or vitamins (V) and amino acids (AA), are essential for the survival and activity of soil bacteria (Swindoll et al. 1988). It has been reported that biostimulation (stimulation of indigenous bacteria) and bioaugmentation (adding bacteria) are performed best in the presence of relative high levels of micronutrients (Ward et al. 1999) (Table 1-5).

Soil properties effecting microbial PAH-degradation in a non-nutritional way

In addition, biodegradation rates are controlled by characteristics of the soil that are not directly involved in bacterial nutrition. The availability of oxygen (O2) and redox- potential (Eh), humidity (H2O), acidity (pH), salinity (IS) and temperature (T) play an important role in the survival and activity of soil bacteria (Table 1-5). Control and tuning of these soil parameters such as temperature, pH, moisture or redox-potential is much more difficult. All supplements added to a soil may have serious wanted or unwanted impact on one or more of these factors. Therefore, careful consideration must be taken in determining the quantity and type of nutrients to add so that optimal

IS, pH and Eh conditions are maintained or created.

• Oxygen & Redox-potential

The biodegradation of PAHs is mostly an aerobic process in which molecular oxygen

(O2) functions as final electron acceptor (EA) in energy generating biochemical pathways. Moreover, oxygenases incorporate oxygen atoms (O) in the aromatic ring of the PAHs. The availability of oxygen for PAH-degradation in soil, however, is controlled by many parameters such as the soil type, the water saturation level of the soil, the presence of other metabolisable substrates that can lead to oxygen depletion and the microbial consumption rate. In soil, aerobic degradation can only take place if a minimum of 10% of the pore space is filled with air (Sims et al. 1993; Wilson et al. 1993; Hurst et al. 1996) (Table 1-5). If air fills less then 1% of the pore volume only

- 51 - Chapter 1 anaerobic processes can take place. Low oxygen concentration in soil has been identified as one of the major parameters limiting the biodegradation of petroleum in saturated soil and ground water (Leahy et al. 1990). Oxygen availability is also expressed in the redox-potential, which defines the total terminal electron availability in the environment. The redox-potential affects the oxidation states of hydrogen, carbon, nitrogen, oxygen, sulfur, etc. The redox-potential of submerged sediments may range from –300 mV for anoxic sediments up to +700 mV for highly aerobic sediments. The lack of oxygen in reduced or anaerobic sediments causes obligate and facultative anaerobic microorganisms to utilize other compounds such as nitrate, sulfate or iron ions as electron acceptors. Although aromatic ring reduction and hydrolytic cleavage under anaerobic conditions has been reported for monoaromatic and polyaromatic hydrocarbons, in general, low redox-potentials decrease the rate of PAH-biodegradation (Cerniglia 1984; Ward et al. 1999). For an aerobic environment, the redox-potential must be above +50 mV. Values for Eh of 100 mV to 400 mV generally indicate relative low oxygen concentrations, but are acceptable for aerobic biodegradation (Sims et al. 1993; Barden 1994). High positive Eh values (400 mV to 800 mV) indicate well-aerated conditions that are optimal for aerobic degradation (De Laune 1981). To increase the oxygen amount in the soil it is possible to till or sparge air. In some cases, oxygen releasing compounds such as hydrogen peroxide or magnesium peroxide (e.g. ORC®) can be introduced in the environment (Rau et al. 2001). Also the supplemental nutrients may react with the compounds in the subsurface or provide an alternate TEA, thus changing the Eh.

• Moisture

Soil moisture is essential to biodegradation since the majority of microorganisms live in the water film surrounding soil particles. Biological degradation processes can only take place if sufficient water is present to keep the soil microorganisms active.

Moisture is for soils generally expressed in terms of ‘water activity’ (aw) or ‘water holding capacity’. The humidity of soil regulates the equilibrium between the water- and gas phase within the pores of the soil matrix and many other physical and chemical soil processes like transport, adsorption, complexation of nutrients, etc.. Tests have shown that a humidity of 70-90% water holding capacity produces optimal

- 52 - Literature Review metabolic rates with a variety of organic pollutants (Hurst et al. 1996; Carmichael et al. 1997) (Table 1-5). A soil humidity range of 70-90% of the water holding capacity is also reported to be optimal for oil and PAH-biodegradation (Sims et al. 1993; Wilson et al. 1993; Barden 1994). Higher water contents is probably one of the reasons why slurry reactors perform better then dry solid reactors or in situ treatments for PAH-degradation. Nutrients and PAHs are more dissolved and more available for bacterial uptake when more water is present. However, in these systems an appropriate mixing or aeration system is crucial since in 90% humidity most of the soil pores are filled with water so that oxygen transfer can become the limiting factor for optimal degradation. In situ, irrigation may be needed to achieve the optimal moisture level (Vidali 2001).

• Acidity

Soil pH is an indicator of hydrogen ion activity in the soil. The pH plays a major role in many physical en chemical soil processes. Since PAHs do not posses acidic or basic groups in their elemental structure, the pH will not directly affect the PAHs. Nevertheless, the behavior of PAHs in soil can be seriously affected by the pH. If the pH rises, humic acids in the soil become more negatively charged, they will loosen their 3-dimensional structure and the sorption of the PAHs will diminish (Chiou et al. 1979; De Laune 1981; Thurman 1985; Murphy 1994; Bastiaens 1998). Most soil organisms can live with pH ranging from 5.5 to 8.5. A pH in the range of 5 to 9 is generally acceptable for biodegradation; a pH of 6.5 to 8.5 is considered optimal (Cookson 1995). In specific, a pH range of 7.0-7.8 was suggested as optimal for microbial PAH-degradation (Wilson et al. 1993). Low pH could negatively affect bacterial growth and thus degradation activity (Baath 1996; Kästner 1998; Alden 2001). Soil pH also affects the availability of other inorganic nutrients. The solubility of phosphorus is maximized at a pH value of 6.5 (Barden 1994). If a soil is too acidic, it is possible to adjust the pH by adding lime (e.g. CaCO3). Neutralization of soil is generally discussed to be favorable for the biodegradation of mineral oil components and PAHs (Leahy et al. 1990; Shen et al. 1996) (Table 1-5).

- 53 - Chapter 1

• Salinity

Salinity is mostly expressed in terms of ‘ionic strength’ (I). The ionic strength refers to the total concentration of ions in solution. The contribution of ions to the ionic strength of the solution increases with the charge of the ion (Formula 1-1). The ionic strength is usually measured as electrical conductivity and calculated via a simplified linear relationship (Formula 1-2).

FORMULA 1-1

2 I = ½ (Σci*Zi ) with ci the molar concentration of ion i

Zi the charge of ion i

FORMULA 1-2

I = 0.000016 * conductivity with conductivity in µS/cm or µmhos/cm

The ionic strength is important for bacteria as it effects electrostatic (Coulombic) interactions between molecules in many biochemical reactions. Proteins have charged surfaces and electrostatic forces determine the interaction of the proteins with their substrates. High salt concentrations such as 0.1 to 0.5 M KCl (i.e. I of 0.1-0.5) can cause dissociation of bacterial membrane proteins (Alden 2001). For bacterial cells the solution may not exceed the maximum salt concentration of 4% w/v NaCl, i.e., I of about 0.700 (Wilson et al. 2003). Experimentally it has been shown that increasing the I of the aqueous solution also increases the extent of bacterial sorption to a variety of natural and artificial surfaces, and enhances the bacterial retention in sand columns in transport experiments (van Loosdrecht et al. 1990a; van Loosdrecht et al. 1990b; Fontes 1991; Gannon 1991; Mills 1994). In addition, the salinity of the groundwater can seriously influence the solubility and the sorption of PAHs to soil-particles (Shiaris 1989). The solubility of naphthalene, phenanthrene and benzo(a)pyrene decreases with increasing I (Eganhouse et al. 1976). In addition, an increase in I also leads to an increase of the sorption-coefficients of PAHs to organic material (Karickhoff et al. 1979). In case of a high I of the soil solution, the negatively charged groups of the humic acids will be shielded by salt ions what allows increased

- 54 - Literature Review absorption of hydrophobic components such as PAHs to the organic particles (Engebretson et al. 1994). In general, for biological processes it is favorable that the soil solution does not exceed an electrical conductivity of 2000µS/cm, i.e., an I of approximately 0,032 (Barden 1994) (Table 1-5). The negative effect of high salinity on the PAH-degradation activity of soil bacteria has been reported several times in estuarine sediments (Kerr et al. 1988; Shiaris 1989) or inoculated soil (Kästner 1998).

• Temperature

Temperature directly affects the rate at which PAH are degraded by microorganisms in natural ecosystems. The soil temperature influenced the biodegradation of organic pollutants by affecting (i) the chemical and physical properties of hydrocarbons, (ii) the rate of metabolic processes in microorganisms and (iii) the composition of the microbial community (Leahy et al. 1990). Seasonal fluxes in heterotrophic activity and PAH degradation rates have been detected several times with the highest activities in summer (20°-25°C) and the lowest activities in winter (Shiaris 1989; Barden 1994). It is unclear whether temperature-related differences in hydrocarbon degradation result from seasonal selection of psychrophilic or mesophilic hydrocarbon-degrading microorganisms or from low-temperature suppression of the degradative capacity of stable PAH-degrading microbial populations that persist during the year. In general, the biodegradation rate increases with increasing soil temperature, and most soil microorganisms show highest activity between 20 and 30°C (Cerniglia 1984; Bauer et al. 1985; Wilson et al. 1993) (Table 1-5). Naphthalene, phenanthrene or anthracene mineralization was not detected at extremes incubation temperatures of 5 and 45°C (Cerniglia 1984; Bauer et al. 1985). It has been shown that the Pseudomonas PAH-catabolic systems located on plasmids are thermo- sensitive as the catabolic plasmids were eliminated at 41-42 °C. Some of these catabolic plasmids had even an inhibition effect on growth of Pseudomonas strains at an elevated temperature, as plasmid free mutants could grow much better on elevated temperature (Kochetkov et al. 1983). For on site and in situ techniques, plastic covering can be used to enhance solar warming in late spring, summer and autumn (Vidali 2001).

- 55 - Chapter 1

• Mineralogy & granularity

The mineralogical composition of soils can have a great influence on the bioavailability of PAHs. PAHs sorb strongly to humic acids or clay (Ortega-Calvo et al. 1998). In addition, mineralogy can also affect the availability of other nutrients such as nitrogen and phosphorus as discussed previously. Soil structure controls the effective delivery of air, water and nutrients. Hence soils with low permeability may not be appropriate for in situ clean-up techniques. To improve soil structure in an ex situ approach, materials such as gypsum or organic matter can be added (Vidali 2001).

• Presence of co-contaminants

Most successful PAH-biodegradation experiments in soil are on lab-scale and use an artificial contaminated soil polluted with one or a selected group PAHs and a selective group of soil bacteria added to the soil. In reality, PAH-contaminated soils mostly contain a complex mixture of pollutants that could interact with each other and with the bacteria degrading the PAHs. PAH contamination always includes a mixture of many PAHs. The quick and complete degradation of mixture of PAHs always require a consortium of different bacteria mineralizing different PAHs. The degradation process of one PAHs by one specific bacterium can be, however, seriously inhibited by the presence of another PAH compound (Bauer et al. 1985; Bouchez et al. 1995), although this inhibition effect was less significant in soil (Bastiaens 1998). A strong similarity in chemical structure between PAHs allows interactions on many different steps in the degradation process, i.e., competition for active zones of the degradation enzymes, accumulation of toxic waste products or interaction on the level of enzyme induction. Heavy metals can be additional pollutants in PAH-contaminated soils. Such heavy metals could disrupt biodegradation processes (Springael et al. 1994). Some PAH-degrading bacteria have combined PAH-degradation capacities with resistance to heavy metals (Kozlova et al. 1999). Also cyanides are very frequent present in PAH-contaminated soils from for example coal gasification plants.

- 56 - Literature Review

• Soil microflora

Other microorganisms present in the soil could have a deleterious effect on PAH- degrading community (Goldstein et al. 1985). They could produce natural antimicrobial substances such as antibiotics or toxins. In addition, bacteriophages and protozoa are wide spread in the environment and could attack or graze on degrading bacteria (Ramadan 1990; Leser et al. 1995).

TABLE 1-5 ENVIRONMENTAL CONDITIONS AFFECTING PAH-DEGRADATION IN SOIL

Conditions required for Optimum values for Parameter References general microbial activity Oil & PAH-degradation (Paul et al. 1989) C/N/P (Wilson et al. 1993) 120/19/4 [mg] (Bouchez et al. 1995) Nutrient content N and P for microbial growth 120/10/1 [mg] (Cookson 1995) 100/15/3 [mg] (Breedveld et al. 2000) 100/10/1 [mg] (Vidali 2001) (Wilson et al. 1993) Aerobic degradation process: 10-40% O Oxygen content 2 (Vidali 2001) minimum air filled pore space of 10% Buried lifts of bed bioreactor 2% to 5% (Hurst et al. 1996) Aerobes and facultative aerobes: (Sims et al. 1993) Redox potential 50 mV > RP < 500 mV 50 mV > RP < 500 mV (Barden 1994) (Cookson 1995) Soil pH 5.5-8.8 6.5-8.0 (Wilson et al. 1993) (Vidali 2001) (Cerniglia 1984) Temperature 15-45 °C 20-37°C (Bauer et al. 1985) (Vidali 2001) IS 0 - 0,056 < 0,032 (Barden 1994) (Sims et al. 1993) Soil moisture 25-85% of water-holding capacity 70-90% (Barden 1994) (Vidali 2001) Contaminants Not to toxic 5-10% oil or PAHs of dry weight of soil (Vidali 2001) Heavy metals Total content < 2000 ppm Total content < 700 ppm (Vidali 2001) Type of soil Low clay or silt content Low clay or silt content (Vidali 2001)

Conclusions

Microbial degradation is considered to be the major route through which PAHs are removed from contaminated environments and therefore a feasible remediation technology. Currently, in situ and ex situ bioremediation techniques are, however, still very inefficient for removal of PAHs from contaminated soil. Two main problems, however often block the efficient biological treatment of PAH-contaminated soil: (i) the limited bioavailability of these compounds due their hydrophobicity and (ii) the

- 57 - Chapter 1 natural soil conditions that reduce the survival and the activity of PAH-degrading bacteria.

Bioremediation is a very soil and site-specific process. If the soil shows no or little natural degrading activity, a consortium of PAH-degrading and nutrients could be added in order to boost bioremediation. The successful implementation of one nutrient-bacteria combination at one site for one contaminant, however, does not guarantee success when applied to another soil type. Currently, there are no specific methods for determining the exact nutrient sources and working conditions one should utilize at a polluted site soil to stimulate bacterial activity. Little practical information about the specific needs of PAH-degrading microorganisms in the soil environment is available. Some studies have reported some possible nutrition and environmental requirements for optimal bacterial activity, but only few could draw conclusions towards final field applications. Moreover, non of these studies were specifically directed towards bacteria specialized in PAH-degradation like Mycobacterium spp. and Sphingomonas spp.. In order to successfully stimulate and use PAH-degrading strains for bioaugmentation, it is crucial further research reveals the true specific macro-, micro- and even trace nutrient requirements of these soil bacteria specialized in PAH-degradation. Especially, since application of nutrient supplements on full- scale includes considerable expenses. One challenge is to learn how microorganisms cope with complex mixtures of PAHs and organic substrates in soil and what occurs when mixed groups of microorganisms are put to work at degrading single or mixed contaminants. Secondly understanding the effects of nutrient type and quantity may enable to develop strategies to change inhibitory situations for the better. So, it will become possible to draw comparisons across different sites dependent upon the site- specific soil characteristics such as nutrient prevalence and other hydro-geological characteristics. In addition, a better understanding of the nutritional needs and degradation kinetics will be useful for the development of mathematical models for carrying out general predictive analysis for bioremediation problems.

There are also important engineering constraints affecting practically the entire field of bioremediation such as problems of how to deliver nutrients, the nature of particular sites, the delivery of organisms, the configuration of reactor systems, and cost and time considerations.

- 58 - Literature Review

Many different environmental and antropogeneous factors influence the survival and activity of PAH-degrading bacteria in the soil. As any other technology engineered biological remediation of PAH-contaminated sites can only become successful if the active system is known and relative controllable. This requires that the degrading soil microorganisms are identified, promoted in the creation and maintenance of an active biomass and directed towards optimal PAH-biodegradation.

- 59 -

“We have a habit in writing articles published in scientific journals to make the work as finished as possible, to cover up all the tracks, to not worry about the blind alleys or describe how you had the wrong idea first, and so on. So there isn't any place to publish, in a dignified manner, what you actually did in order to get to do the work.”

- Richard Phillips Feynman (1918-1988) -

- 60 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

CHAPTER 2

OCCURRENCE AND DIVERSITY OF FAST-GROWING MYCOBACTERIUM SPECIES IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHs)*

* REDRAFTED AFTER: LEYS NATALIE, RYNGAERT ANNEMIE, BASTIAENS LEEN, WATTIAU PIERRE, TOP

EVA, VERSTRAETE WILLY, SPRINGAEL DIRK (REVISED) OCCURRENCE AND DIVERSITY OF FAST-

GROWING MYCOBACTERIUM SPECIES IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC

HYDROCARBONS (PAHS), FEMS MICROBIOL. ECOL.

ABSTRACT

Mycobacterium species using polycyclic aromatic hydrocarbons (PAH) as sole source of carbon and energy may be essential members of the bacterial populations degrading PAHs in the environment. To study the natural role and diversity of PAH- degrading Mycobacterium communities in contaminated soils, a culture-independent fingerprinting method based on PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) was developed. As so far all PAH-degrading Mycobacterium isolates could be placed in the phylogenetic branch of the ‘fast-growing’ Mycobacterium species, new PCR primers were selected to specifically target 16S rRNA genes of fast-growing Mycobacterium species. The new primer set proved to be highly selective for the target group in PCR and single-band DGGE profiles were obtained for most Mycobacterium strains tested. Strains belonging to the same species had identical DGGE fingerprints, and in most cases but not all, these fingerprints were typical for one species, allowing partial differentiation between species in a Mycobacterium population. Mycobacterium strains inoculated in soil were detected with a detection limit of 106 CFU g-1 of soil using the new primer set alone or of circa 102 CFU g-1 in a nested PCR approach combining eubacterial and the new Mycobacterium specific primers. The PCR-DGGE detection method was used to rapidly assess the Mycobacterium population structure of several PAH-contaminated soils of diverse origin and different overall contamination profiles, pollution concentrations and chemical-physical soil characteristics. In most PAH-contaminated soils well-known PAH-degrading species like M. frederiksbergense and M.

- 61 - Chapter 2 austroafricanum were detected. Interestingly, 16S rRNA genes related to M. tusciae sequences, a Mycobacterium species so far not reported in relation to biodegradation of PAHs, were detected in all soils.

INTRODUCTION

Polycyclic aromatic hydrocarbons (PAHs) are hazardous environmental pollutants that are found in high concentrations in the surface soil of old gas factories and wood preservation plants (Cerniglia 1992). In spite of the limited bioavailability and poor biodegradability of PAHs, different bacteria, often Mycobacterium strains, have been isolated that are able to use PAHs as sole source of carbon and energy (Guerin et al. 1988; Bastiaens 1998; Churchill et al. 1999; Bastiaens et al. 2000; Solano-Serena et al. 2000; Willumsen et al. 2001a). So far, all PAH-biodegrading Mycobacterium isolates (Guerin et al. 1988; Briglia et al. 1994; Bastiaens 1998; Churchill et al. 1999; Poelarends et al. 1999; Yagi et al. 1999; Bastiaens et al. 2000; Schrader et al. 2000; Solano-Serena et al. 2000; Willumsen et al. 2001a), have been placed in the phylogenetic branch of the ‘fast-growing Mycobacterium species’. In the Mycobacterium genus phylogenetic tree, the ‘fast-growing Mycobacterium species’ form a coherent line of descent, distinct from the more recently evolved slow-growers within which the overt pathogens are clustered (Rogall et al. 1990; Stalh et al. 1990; Pitulle et al. 1992; Tortoli 2003). The ‘fast-growing Mycobacterium species’ are a group of Mycobacterium strains, mostly of environmental origin, that are, based on growth and biochemical characteristics and infectious properties (i.e. Mycobacterium species of Bio Safety Level 1, growth within 7 days) very different from pathogenic and facultative pathogenic more slowly growing species like M. avium, M. tuberculosis, M. leprae or M. ulcerans (i.e. Mycobacterium species of Bio safety level 2 & 3, growth after more than 7 days). The diversity of fast-growing Mycobacterium species in the environment is still greatly unknown, but could be of major interest for bioremediation of PAH- contaminated soils. Therefore, methods for community analysis and monitoring of indigenous and/or inoculated fast-growing PAH-degrading Mycobacterium strains in soil are needed. Direct culture-independent detection methods are more preferable above indirect culture-dependent techniques for detection of Mycobacterium strains because (i) a large fraction of cells in soil is hard to culture or even believed to be

- 62 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils. unculturable (Viable But Non Culturable state) (Staley et al. 1985; Amann et al. 1995; Ghezzi et al. 1999), (ii) the hydrophobic Mycobacterium cells are known to adhere strongly to organic soil particles resulting in their difficult recovery (Barry et al. 1998; Draper 1998), and (iii) Mycobacterium species, even the ‘fast-growers’, are relatively slow-growing organisms in comparison to other soil bacteria, which makes them easily overgrown by other bacteria in culture media (Allen 1998). In addition, molecular PCR-based methods had proven to be successful for the diagnosis of Mycobacterium diseases in humans (Böddinghaus et al. 1990; De Beenhouwer et al. 1995; Kox et al. 1995; Kox et al. 1997; van der Heijden et al. 1999) and fish (Talaat et al. 1997) and for the identification of environmental infection sources of Mycobacterium opportunistic pathogens such as M. avium and M. ulcerans in plants, water and soil (Schwartz et al. 1998; Mendum et al. 2000; Stinear et al. 2000). PCR amplification of variable 16S rRNA gene-fragments combined with direct analysis of amplicons by Denaturing Gradient Gel Electrophoresis (DGGE) is a commonly used technique for rapid molecular assessment of the community diversity. In all previous studies, however, the PCR primers were designed to reveal the presence of slow- growing Mycobacterium species in the tested samples and were never combined with a method for direct community diversity analysis like DGGE. Moreover, most importantly, none of the described primer sets were specific enough to target preferentially fast-growing Mycobacterium species, belonging to non-pathogenic and non-opportunistic species. We describe in this study the development of a new set of non-degenerated primers that annealed as exclusively as possible to 16S rRNA genes of fast-growing Mycobacterium strains and that amplified a short fragment suited for DGGE a culture- independent method for fingerprinting of only the fast-growing Mycobacterium species. The PCR-DGGE method was theoretically and practically evaluated for detection of fast-growing Mycobacterium species in soil samples and applied to examine the Mycobacterium diversity in PAH-contaminated soils.

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MATERIALS AND METHODS

Bacterial strains and growth conditions. The bacterial strains used in this study are described in Table 2-1. For DNA-extraction purposes, strains other than Mycobacterium strains were cultivated in 869-broth (Mergeay et al. 1985) while Mycobacterium strains were cultivated in Middelbrook 7H9 Broth (DIFCO, Kansas City, USA). For inoculation purposes, Mycobacterium strains were cultivated in a phosphate buffered minimal liquid medium previously described (Wick et al. 2001), containing 2 g l-1 of anthracene or pyrene crystals (ACROS Organics, Fisher Scientific, Boston, USA) floating in the medium as the sole carbon and energy source. All cultures were grown in the dark on an orbital horizontal shaker at 200 rpm at a constant temperature of 30 °C.

TABLE 2-1 BACTERIAL STRAINS USED IN THIS STUDY

Reported pollutant 16S rRNA gene Organism Reference or origin degrading capacity Accession N° PHYLUM OF HIGH G+C GRAM POSITIVE BACTERIA Order of Actinomycetales Actinomyces sp. A1008 NR NR (Heuer et al. 1997) Actinosynnema mirum 101 NR X84447 DSM 43827T Arthrobacter sulfureus 8-3 NR X83409 DSM 20167T Kineospora aurantiaca A/10312 NR X87110 DSM 43858T Microbispora rosea IMRU37485 NR NR ATCC 21946 Micromonosprora chalcea A0919 NR NR (Heuer et al. 1997) Micromonosprora chalcea A2868 NR NR (Heuer et al. 1997) Micromonosprora chalcea A2894 NR NR (Heuer et al. 1997) Planomonospora parontospora B-987 NR AB028653 DSM 43869T Promicromonospora citrea INMI 18 NR X83809 DSM 43110T Streptomyces albus A0818 NR NR (Heuer et al. 1997) Streptomyces albus A1893 NR NR (Heuer et al. 1997) Streptomyces albus A2198 NR NR (Heuer et al. 1997) Streptomyces albus A3986 NR NR (Heuer et al. 1997) Streptomyces aureofaciens A-377 NR NR DSM 40127T Streptomyces rutgersensis BJ-608 NR NR DSM 40830T Streptomyces phaeofaciens T-23 NR D44381 DSM 40367T Streptosporangium album A0958 NR NR (Heuer et al. 1997) Suborder Corynebacterineae Dietziaceae family Dietzia maris VM0283 diesel NR (Wattiau et al. 1999) Dietzia maris IMV 195 NR X79290 DSM 43627T Corynebacteriaceae family Corynebacterium glutamicum 2247 NR NR DSM 20411 Tsukamurallaceae family Tsukamurella paurometabola NR AF283280 DSM 20162T Nocardiaceae family Nocardia asteroides N3 NR NR (Heuer et al. 1997) Nocardia coeliaca AB.4.1.b NR NR DSM 44595T Pseudonocardia hydrocarbonoxydans NR X76955 DSM 43281T Rhodococcus erythropolis ICPB 4417 NR X81929 DSM 43066T Gordoniaceae family Gordonia hydrophobica 1610/1b NR X87340 DSM 44015T Gordonia amarae Se 6 NR X80601 DSM 43392T Mycobacterinaceae family Mycobacterium aichiense 5545 NR X55598 DSM 44147T Mycobacterium alvei CR-21 NR AF023664 DSM 44176T Mycobacterium aurum 358 NR X55595 DSM 43999T Mycobacterium vanbaalenii PYR-1 nap, phe, fan, pyr U30662 DSM 7251 T Mycobacterium austroafricanum E9789 NR X93182 DSM 44191T Mycobacterium austroafricanum VM0456 phe AF44622 (Springael et al. unpublished) Mycobacterium austroafricanum VM0450 phe AF44623 (Springael et al. unpublished) Mycobacterium austroafricanum VM0451 phe, pyr, fan, ant AF44624 (Springael et al. unpublished) Mycobacterium austroafricanum VM0447 phe AF44625 (Springael et al. unpublished) Mycobacterium austroafricanum VM0452 phe, fan AF44626 (Springael et al. unpublished) Mycobacterium austroafricanum VM0573 phe AF44627 (Springael et al. unpublished) Mycobacterium chlorophenolicum PCP-1 PCP X79094 DSM 43826T

- 64 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

Mycobacterium diernhoferi SN1418 NR X55593 DSM 43524T Mycobacterium frederiksbergense FAn9 fan, phe, pyr AJ276274 DSM 44346T Mycobacterium frederiksbergense LB501T ant, phe, diesel AJ245702 (Bastiaens et al. 2000) Mycobacterium frederiksbergense VM0503 fan AF44628 (Springael et al. unpublished) Mycobacterium frederiksbergense VM0531 phe, pyr AF44629 (Springael et al. unpublished) Mycobacterium frederiksbergense VM0458 phe AF44630 (Springael et al. unpublished) Mycobacterium frederiksbergense VM0579 phe AF44631 (Springael et al. unpublished) Mycobacterium frederiksbergense VM0585 phe, fan AF44632 (Springael et al. unpublished) Mycobacterium gilvum SM35 NR X81996 DSM 44503T Mycobacterium gilvum BB1 phe, flu, pyr, fan X81891 DSM 9487 Mycobacterium gilvum HE5 mor, prl, pip AJ012738 DSM 44238 Mycobacterium gilvum LB307T phe, pyr, dib, fan, diesel AJ245703 (Bastiaens et al. 2000) Mycobacterium gilvum VM0505 ant AF44633 (Springael et al. unpublished) Mycobacterium gilvum VM0504 ant AF44634 (Springael et al. unpublished) Mycobacterium gilvum VM0552 phe, pyr AF44635 (Springael et al. unpublished) Mycobacterium gilvum VM0442 phe, fan, ant AF44636 (Springael et al. unpublished) Mycobacterium gilvum LB208 phe, pyr, fan, diesel AJ245704 (Bastiaens et al. 2000) Mycobacterium gilvum VM0583 ant AF44637 (Springael et al. unpublished) Mycobacterium hodleri EM12 fan X93184 DSM 44183T Mycobacterium komossense Ko2 NR X55591 DSM 44078T Mycobacterium neoaurum 3503 NR M29564 DSM 44074T Mycobacterium parafortuitum 311 NR X93183 DSM 43528T Mycobacterium peregrinum 6020 NR AF058712 DSM 43271T Mycobacterium petroleophilum RF002 fan, phe, pyr U90876 (Lloyd-Jones et al. unpublished) Mycobacterium vaccae VM0587 fan AF44638 (Springael et al. unpublished) Mycobacterium vaccae VM0588 fan AF44639 (Springael et al. unpublished) Mycobacterium sp. WF2 fan U90877 (Lloyd-Jones et al. unpublished) Mycobacterium sp. GP1 DBA AJ012626 (Poelarends et al. 1999) PHYLUM OF FLAVOBACTERIA Flavobacterium resinovorum oleoresins NR ATCC 33545T PHYLUM OF PROTEOBACTERIA α-subdivision Agrobacterium luteum A61 NR NR DSM 5889T Brevundimonas diminuta 342 NR AJ227778 DSM 7234T Sphingomonas chlorophenolica PCP X87161 DSM 7098T β-subdivision Ralstonia metallidurans CH34 NR Y10824 DSM 2839T Burkholderia sp. JS150 benzene derivates AF262932 DSM 8530 γ-subdivision Aeromonas enteropelogenes J11 NR X71121 DSM 6394T Acinetobacter calcoaceticus 46 NR AJ247199 DSM 30006T Pseudomonas putida nap, phe, flu, fan NR DSM 8368 δ-subdivision Desulfobacter latus AcRS2 NR AJ441315 DSM 3381T Desulfonema magnum 4be13 NR U45989 DSM 2077T Desulfobulbus rhabdoformis M16 NR U12253 DSM 8777T T = species type strain; NR = Not Reported; nap = naphthalene; fan = fluoranthene; pyr = pyrene; flu = fluorene; phe = phenanthrene; ant = anthracene; dib= dibenzothiophene; mor = morpholine; prl = pyrrolidine; pip = piperidine; PCP = pentachlorophenol; DBA = 1,2-di-bromoethane.

Soils used in this study. Soil samples were taken from different anthropogenic PAH-contaminated sites (Table 2-2). The soil texture, pH (DIN Method 38414, S4), total carbon content (TC), total inorganic carbon (TIC) content (hydrolysis method) and total organic carbon (TOC) content of each soil sample was determined (ISO-CEN EN Method 1484). The soils were chemically analyzed for the 16 PAHs legislated by the U.S. Environmental Protection Agency. PAHs were extracted through an Accelerated Solvent Extraction (ASE 200 Accelerated Solvent Extractor, Dionex Corp., Sunnyval, CA, USA) (EPA Method 3545). ASE-extracts were purified over an internal silica phase in the extraction cell (i.e. in thimble clean up) followed by an alumina column. Quantification was done by capillary gas chromatography (Carlo Erba MFC 500 with split/splitless injector) coupled to a mass- spectrophotometric detector (quadrupole-type, Fisons QMD 100) (EPA Method 8270). The total concentration of mineral oil present in the soil sample was determined after an ultrasonic tetrachloroethene extraction followed by a FLORISIL clean up (U.S. Silica Company, Berkeley Springs, USA) using an infrared quantification at 2925, 2958 and 3030 cm-1 (NEN Method 5733).

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TABLE 2-2 SOIL SAMPLES USED IN THIS STUDY

Soil TOC PAH conc. Oil conc. DNA conc.* Soil Origin pH MYCO‡ Nested † type (%) (mg kg-1) (mg kg-1) (µg g-1) S587 Corn field (Belgium) sand 5.5 2.15 0.289 < 50 31.00 + ND S588 Horse pasture (Belgium) sand 6.0 2.46 0.391 < 50 18.00 + ND S585 Pine tree forest (Belgium) sand 5.8 3.19 0.673 < 50 31.75 + ND S589 Ditch in agricultural area (Belgium) sand 5.8 4.24 0.721 < 50 49.50 + ND S592 Vegetable garden (Belgium) sand 7.0 3.16 1.011 < 50 38.25 + ND S584 Compost heap (Belgium) sand 7.3 7.04 1.063 < 50 27.75 + ND S591 Non-paved land road (Belgium) sand 9.0 0.76 3.357 < 50 6.25 NP ND TB3 Coal gasification plant (Belgium) sand 8.23 1.52 14 < 50 2.65 + + K3840 Gasoline station site (Denmark) Sand 8.20 0.50 20 98 2.75 + + B101 Coal gasification plant (Belgium) Sand 7.00 2.63 107 70 27.25 + + E6068 Gasoline station site (Denmark) Sand 7.96 9.94 258 300 5.40 + + TM Coal gasification plant (Belgium) Sand 8.00 3.85 506 4600 4.75 + + Barl Coal gasification plant (Germany) Gravel 8.90 4.63 1029 109 6.15 NP NP AndE Railway station site (Spain) Clay 8.10 2.35 3022 2700 3.40 NP + * DNA recovery per g soil, mean value of 2 parallel extractions of 1 gr of soil. ‡ result of direct PCR with Mycobacterium specific primers MYCO66f and GC40-MYCO600r on soil DNA extract: + = detectable PCR product, NP = no detectable PCR product, ND = not determined. † result of nested PCR with eubacterial primers 27f and 1492r followed by Mycobacterium specific primers MYCO66f and GC40-MYCO600r on soil DNA extract: + = detectable PCR product, NP = no detectable PCR product, ND = not determined.

Design of 16S rRNA gene primer set specific for fast-growing Mycobacterium strains. Primer sequences were selected from a multiple alignment of circa 200 16S rRNA genes (Genbank, NCBI) of circa 100 fast- and 100 slow-growing Mycobacterium species constructed with the Bionumerics software (Applied Maths, Version 2.50). The alignment was further analyzed with the PLOTCON program (EMBOSS, Version 1.9.1) to identify conserved and variable gene regions. Based on the alignment, the new Mycobacterium specific primers were selected in gene regions that are conserved within the group of fast-growing Mycobacterium species but as variable as possible within slow- growing Mycobacterium species. In addition, for optimal species differentiation in DGGE-analysis of the PCR-products (see below), the primers were selected so that they amplified a region between 200 bp and 600 bp long with high variability. The selectivity of the selected primers was evaluated by visual analysis of the primer region within the constructed alignment of Mycobacterium rrn genes, by the Sequence Match program (RDP II) (Cole et al. 2003) and by the Advanced Blast Search program (Genbank, NCBI) (Altschul et al. 1990). The best primer combination consisted of forward primer MYCO66f (5'-CATGCAAGTCGAACGGAAA-3', E. coli position 66 to 84) and reverse primer MYCO600r (5'-TGTGAGTTTTCACGAACA-3', E. coli position 600 to 583). A 40 basepair long GC- clamp (Muyzer et al. 1993; Muyzer et al. 1996) was attached to the 5' end of the MYCO600r primer for DGGE analysis of the Mycobacterium amplicons. This new primer couple MYCO66f and GC40- MYCO600r amplified a 538 bp sequence of the 16S rRNA gene resulting in a PCR-product of 578 bp. DNA-extraction. Genomic DNA from pure bacterial cultures was obtained as described by Belisle et al. (Belisle et al. 1998). The DNA recovery was approximately 2.7 to 27.3 µg DNA g-1 soil. For PCR purposes, the DNA-concentration was adjusted to a final concentration of 100 ng µl-1. For fast-growing Mycobacterium strains, 100 ng of template DNA corresponds to 1.2-1.9×107 cell equivalents of genomic DNA and 2.4-3.8×107 copies of PCR targets assuming a genomic molecular weight of 3.13- 5.20×109 Daltons per cell and two 16S rRNA gene copies per genome (Bercovier et al. 1986). DNA was extracted from 1 g soil using a protocol described by Boon et al. (Boon et al. 2000). After

- 66 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils. purification over a Wizard column, the DNA concentration in the 50µl soil extract was measured photospectroscopically. To assure that the soil DNA was of PCR quality, dilution series of soil DNA extracts were tested by PCR with universal eubacterial 16S rRNA gene primer pair GC40-63f and 518r as previously described (Marchesi et al. 1998). PCR amplification of pure strain and soil DNA. The PCR protocol used with the MYCO66f and MYCO600r primer pair consisted of a short denaturation of 15 s at 95°C, followed by 50 cycles of denaturation for 3 s at 94°C, annealing for 10 s at 50°C, elongation for 30 s at 74°C, and a final extension for 2 min at 74°C. PCR was performed on Biometra or PerkinElmer Thermalcylcers. PCR mixtures contained 100 ng of pure strain DNA or dilutions of soil DNA as templates, 1 U Taq polymerase, 25 pmol of the forward primer, 25 pmol of the reverse primer, 10 nmol of each dNTP, and 1 × PCR buffer in a final volume of 50 µl. Primers designed by Cheung and Kinkle (MycF and MycR) (Cheung et al. 2001) were used in PCR as described. All primers were synthesized by Westburg (Westburg BV, Leusden, The Netherlands). The Taq polymerase, dNTPs and PCR buffer were purchased from TaKaRa (TaKaRa Ex TaqTM, TaKaRa Shuzo Co., Ltd., Biomedical Group, Japan). DGGE analysis. The PCR-products were examined on 1.5 % agarose gels (MetaPhor, BioWhittaker, Labtrade Inc., Miami, Florida, USA) and directly used for DGGE analysis on polyacrylamide gels as described previously (Muyzer et al. 1998a). Optimal denaturing conditions were defined based on the theoretical melting temperatures of amplification fragments calculated with the Melt Analysis Software (Version 1.0.1, INGENY). A 6 % polyacrylamide gel with a denaturing gradient of 40 % to 75 % (100% denaturant gels contain 7M urea and 40 % formamide) was used for DGGE-apparatus. Electrophoresis was performed at a constant voltage of 130 V for 16 h 40 m in 1 × TAE running buffer at 60 °C in the DGGE-machine (INGENYphorU-2, INGENY International BV, The Netherlands). After electrophoresis, the gels were stained with 1 × SYBR Gold nucleic acid gel stain (Molecular Probes Europe BV, Leiden, The Netherlands) and photographed under U.V. light using a Pharmacia digital camera system with Liscap Image Capture software (Image Master VDS; Liscap Image Capture, Version 1.0, Pharmacia Biotech, Cambridge, England). Photofiles were processed and analyzed with the Bionumerics software (Version 2.50, Applied Maths, Kortrijk, Belgium). Sensitivity of PCR-DGGE method. To study the sensitivity of the PCR-DGGE method, a known amount of viable Mycobacterium cells were added to white sand (a model soil matrix) or natural soil samples at different final cell densities (i.e. a 10-fold dilution series of approximately 108 to 101 cells g- 1) prior to DNA-extraction. Cells were harvested from liquid cultures, washed twice and added in 100 µl aqueous suspensions to 1 g of soil. One, two or three different Mycobacterium strain (LB501T, VM0552 and DSM 43524T) were separately or simultaneously added in different cell densities to assess the effect of cell ratios on the detection sensitivity for each single strain within a Mycobacterium population. The total soil DNA was subsequently used in PCR with the MYCO-primers and PCR- products were analyzed by DGGE. Sequence analysis of amplified 16S rRNA gene fragments. PCR products of Mycobacterium 16S rRNA genes were cloned into plasmid vector pCR2.1-TOPO using the TOPO Cloning Kit (N.V. Invitrogen SA, Merelbeke, Belgium) as described by the manufacturer. DGGE patterns of cloned fragments were compared with the fingerprints of the parent soil Mycobacterium community to identify

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which signals from the community fingerprint were cloned. A 500 bp long fragment was sequenced (Westburg BV, Leusden, The Netherlands) from a selection of clone inserts with different DGGE- patterns. The sequences were analyzed with the 'Chimera Check' program (RDPII) (Cole et al. 2003) to detect possible chimeras and with ‘Blast Search’ program (Genbank, NCBI) (Altschul et al. 1990). Cloned sequences were imported into an alignment of Mycobacterium 16S rRNA genes and edited manually to remove nucleotide positions of ambiguous alignment and gaps. Sequence similarities were calculated over the 16S rRNA gene fragment between the MYCO-primers, corrected using Kimura's two-parameter algorithm to compensate for multiple nucleotide exchange and used to construct a distance-based evolutionary tree with the Neighbor-Joining algorithm (Saitou et al. 1987). The topography of the branching order within the dendrogram was evaluated by using the Maximum- Likelihood and the Maximum-Parsimony character-based algorithms in parallel combined with bootstrap analysis with a round of 500 reassemblings. An out-group of the closely related genera Rhodococcus and Dietzia was included to root the tree. Nucleotide sequence accession numbers. The partial 16S rRNA gene sequences of Mycobacterium clones reported in this study are available from GenBank under accession numbers AY148197 to AY148217.

RESULTS

Design of specific primers for fast-growing PAH-degrading Mycobacterium strains. A new specific primer set was designed to amplify the 16S rRNA genes of fast- growing Mycobacterium species. Based on an alignment of approximately 200 sequences, the 16S rRNA genes of fast- and slow-growing Mycobacterium species appeared highly conserved in comparison to other bacteria, i.e., only a few short well- defined regions within the gene were found to be highly variable (data not shown). A minimum similarity of 94% over the total length of the 16S RNA gene was found for all fast-growing Mycobacterium species, making it is very difficult to select strictly group- or species- specific primers. The best possible primer combination was selected from the alignment taking into account the amplicon length and amplicon variability and the Blast and sequence Match results of both primers. The sequence of the forward primer MYCO66f (E. coli locations 66-84) was conserved in 300 rrn gene sequences of mainly fast- but also some slow-growing Mycobacterium strains of the approximately 900 Mycobacterium sequences currently available in the Genbank database (NCBI) (Table 2-3). The MYCO66f primer also

- 68 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils. aligned 100% with Corynebacterium, Phytoplasma, Gordonia and Propionibacterium 16S rRNA genes. The sequence of primer MYCO600r (E. coli locations 600-583), however, was 100% conserved in only 165 sequences of exclusively fast-growing environmental Mycobacterium strains, including all known PAH degrading species (Table 2-3). It clearly differed from most other 16S rRNA gene sequences from slow-growing Mycobacterium strains and non-Mycobacterium strains with 1 to 7 mismatches of the 18bp long primer region (Table 2-3). As for some slow-growing mycobacteria the mismatches with the MYCO660r primer were more concentrated to the 5’ in stead of the 3’ primer end, amplification of the 16S rRNA genes may be possible. Nevertheless, the MYCO660r primer was our best possible choice and showed more mismatches with sequences from slow-growing Mycobacterium than any other primer decribed so far. The high number of mismatches will hinder or prevent the amplification of template from most slow-growing Mycobacterium species and most species not belonging to the Mycobacterium genus. The primer couple MYCO66f and MYCO600r produced products of the appropriate size with the DNA obtained from the 40 tested Mycobacterium strains representing different fast-growing environmental and PAH degrading species (Table 2-1). Due to the risks associated with most slow-growing Mycobacterium species classified as 'Biosafety Level 2 & 3’-agents (Anonymous 1999) only fast-growing reference strains were tested. Nevertheless, PCR reactions conditions were optimized and always performed under very stringent reaction conditions (very short annealing times) to minimize amplification of 16S rRNA genes with only few mismatches located at the 5’ primer end as found in some slow-growing Mycobacterium species. No PCR- products were obtained with DNA from strains belonging to other related genera such as Actinomyces, Arthrobacter, Dietzia, Corynebacterium, Nocardia, Sphingomonas, Burkolderia, Acinetobacter, Desulfobacter and more (Table 2-1). Moreover, non of the cloned sequences obtained from PCR products with soil DNA extracts as template, showed close relationships to any slow-growing Mycobacterium species or species not belonging to the Mycobacterium genus.

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TABLE 2-3 DNA-SEQUENCE HOMOLOGY BETWEEN THE MYCOBACTERIUM GENUS SPECIFIC PRIMERS AND THE 16S RRNA GENE SEQUENCE OF SOME REFERENCE MYCOBACTERIUM SPECIES

16S rRNA Organism Primers† gene * MYCO66f MYCO600r (E.coli 66-84) (E. coli 600-583) 5' - CATGCAAGTCGAACGGAAA - 3' 5' - TGTGAGTTTTCACGAACA - 3' Fast-growing Mycobacterium species M. aichiense X55598 ------M. alvei AF023664 ------M. aurum X55595 ------M. austroafricanum X93182 ------M. chlorophenolicum X79094 ------M. diernhoferi X55593 ------A ------M. frederiksbergense AJ276274 ------M. gilvum X81996 ------M. hodleri X93184 ------M. komossense X55591 ------T ------M. neoaurum M29564 region unsequenced ------M. parafortuitum X93183 ------M. peregrinum AF058712 ------M. petroleophilum RF002 U90876 ------M. vaccae VM0587 AF44638 ------M. sp. AJ012738 ------M. sp. WF2 U90877 ------Slow-growing Mycobacterium species M. gordonae X52923 ------T - - C- - - - - A ------M. genavense X60070 ------CCCCCGA ------M. branderi X82234 ------G - - - M. leprae X55587 ------C - - - - - A ------NN M. tuberculosis H37Rv NC_000962 ------C - - - - - A ------M. ulcerans X88926 ------C - - - - - A ------Non-Mycobacterium bacteria Gordonia terrae AB111113 AF397061 ------C - - A - - - CACA - - CG- Corynebacterium sp. AF262996 ------CCG - TA - - - CACA - - CG- Phytoplasma sp. AF500334 ------T G A- A - - AAACTAG -G- Rhodococcus globerulus X77779 ------T ------A - - - -G- Dietzia maris X79290 ------G - - - T - - C -A - GA - - - - - AG- - - G-

* Accession No of 16S rRNA gene sequence in the Genbank (NCBI, Rockville Pike Bethesda, USA) † N = A or T or G or C, Dashes indicate homologous sequences.

Differentiation of fast-growing Mycobacterium species by DGGE-analysis. In order to examine the potential of the PCR-DGGE method based on the new primer set to differentiate between species within a Mycobacterium community, pure strain DGGE-patterns of a variety of fast-growing Mycobacterium strains were compared. Different Mycobacterium isolates belonging to the same species showed identical DGGE-patterns. This was observed for 8 tested M. austroafricanum related strains, 7 tested M. frederiksbergense related strains and 10 tested M. gilvum related strains (data not shown). Usually different species showed different DGGE fingerprints (Figure 2-1). For example, the PCR-products obtained for the M. frederiksbergense strain (lane 16) migrated clearly differently from the products from M. austroafricanum (lanes 3 & 4) or M. gilvum (lane 9) strains.

- 70 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

1T; ;

2; 1; P DSM44346T; DSM43826T; e DSM7251 DSM4419 RF002; . WF . G DSM43528T; p p

s sp. VM0579; sp. VM0585; s gens DSM44078T; DSM43271T; sp. DSM44238; ilum 0587; DSM44074T; er DSM43524T; m M44183T; itum h DSM44147T. M44503T; i m b nse M iks DS V DS DSM43999T; DSM44176T; urum e grinu er e mosse troleop

ed o cca lorophenolicum k parafortu aichiense alvei fr neoa pe per ...... gilvum hodleri aurum ch austroafricanum va diernhofer austroafricanum ...... M M M M M M. Mycobacterium Mycobacterium M Mycobacterium M M M Mycobacteriu M M M M

, Mycobacterium , M , M 20, 21, 16, 17, 18, 19, 15, 14, 8, 9, 10, 11, 12 13, 7, 2 3 4, 5, 6, 1, * * **

FIGURE 2-1 MYCOBACTERIUM SPECIES DIFFERENTIATION BY DGGE-ANALYSIS OF MYCOBACTERIUM DNA-FRAGMENTS AMPLIFIED WITH THE MYCOBACTERIUM GENUS SPECIFIC PRIMER PAIR MYCO66F AND GC40-MYCO600R. DGGE-fingerprints of strains were compared by Bionumerics software based on co-running standard (not shown). The symbol * indicates multiple band DGGE patterns for single strains with arrows indicating cloned and sequenced bands.

However, in some cases the differences in migration between different species were minor, due to the strong conservation of the Mycobacterium 16S rRNA genes. The amplified rrn gene fragments of species such as M. spaghni (not shown), M. hodleri (lane 8) and M. gilvum (lane 9) with a similarity of 99 to 100% and could not be differentiated by DGGE. For comparison, the amplicons of primer pairs, i.e., MYCO66f and GC40-MYCO600r and MycF and GC40-MycR (Cheung et al. 2001), were analyzed under identical DGGE-conditions. Although a different fragment of the 16S rRNA gene was amplified, PCR-DGGE with both primer sets resulted in the same species differentiation degrees for all tested Mycobacterium strains (data not shown). The species specific DGGE fingerprints usually displayed one single band. However, some strains revealed extra bands in comparison to other members of the same species (Figure 2-1). For example strains DSM 7251 (lane 3) and VM0450 (data not shown) both showed a similar pattern with 5 bands, strains VM0579 (lane 15), VM0531 (data not shown) and VM0503 (data not shown) displayed 3 bands and

- 71 - Chapter 2

VM0585 (lane 14) even 10 bands. The same number of DGGE-bands per strain were obtained when using an other Mycobacterium specific primer the MycF and MycR primers designed by Cheung and Kinkle on the test strains (Cheung et al. 2001) (data not shown). The sequences of these multiple bands for one strain showed only very limited variation, i.e., 98-99% similarity or max. 4 point mutations and deletions over the 500 bp sequenced fragment, and chimera analysis (Cole et al. 2003) of sequenced bands was negative. Multiple random cloning attempts of these PCR products in total or of single excised bands to selectively clone more fainter or higher located additional bands to further investigate their sequence divergence were not successful.

Sensitivity of the Mycobacterium specific PCR-DGGE protocol. To examine the sensitivity of the PCR-DGGE protocol to detect Mycobacterium strains in soil, a known decreasing amount of Mycobacterium sp. LB501T cells were added prior to DNA-extraction to white sand and different PAH-contaminated soil samples (Table 2-2). For some contaminated soils there was a clear inhibitory effect of the soil matrix on the PCR amplification, so the soil DNA template was diluted 1:10 or 1:100 prior to PCR. In addition, white sand was used as a model soil matrix. In a PCR reaction with primer pair MYCO66f and GC40-MYCO600r, LB501T cells could generally be detected until a minimum cell concentration of 106-108 cells per of soil. The same order of detection limit was obtained when using an other Mycobacterium specific primer the MycF and MycR primers designed by Cheung and Kinkle on the same DNA-extracts (Cheung et al. 2001) (data not shown). Similar results were also obtained when two or three different Mycobacterium strains (LB501T and VM0552 or LB501T, VM0552 and DSM 43524T) were simultaneously added to white sand in equal cell concentrations (ratios 1:1 or 1:1:1 ratio), i.e., all strains were detected equally well until a concentration of 106 to 107 CFU g-1 soil (Figure 2-2). Attempts to improve significantly the detection limit by optimizing the DNA extraction and purification protocol or reducing length of the GC-clamp were not successful (data not shown). Only via a nested PCR approach, combining a first PCR with universal eubacterial primers and a second PCR with the MYCO-primers, the detection limit could be lowered to circa 102 cells per gram of soil (data not shown).

- 72 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

To assess the impact of unequal cell concentration ratios on the detection sensitivity, different concentrations of VM0552 or VM0552 and DSM 43524T (1:1) cells were added to white sand in the presence of a constant concentration of circa 108 CFU g-1 of LLB501T. Declining cell amounts of VM0552 and DSM 43524T could be detected in the presence of 108 CFU g-1 LB501T cells until a concentration of 106 CFU g-1 (Figure 2-2).

A VM0552: LB501T in 1:1 DSM43524: VM0552:LB501T in 1:1:1

8 6 4 2 7 5 3 1 10 10 10 10 10 10 10 10 * *

LB501T VM0552

DSM43524

B VM055 2:LB501T DSM43524: VM0552:LB501T

1:1 1:2 1:4 1:10 1:100 1:10000 1:1:1 1:1:2 1:1:4 1:1:10 1:1:100 1:1:10000 * 8 8 8 8 *8 8 7 7 7 7 7 7 10 10 10 10 10 10 10 10 10 10 10 10 LB501T VM0552 108 5x107 2x107 107 106 104 107 5x106 2x106 106 105 103 DSM43524 107 5x106 2x106 106 105 103

FIGURE 2-2 DETECTION EFFICIENCY OF THE PCR-DGGE METHOD USING MYCOBACTERIUM GENUS SPECIFIC PRIMER PAIR MYCO66F AND GC40-MYCO600R (A) PCR-DGGE detection of the simultaneously added M. frederiksbergense LB501T, M. gilvum VM0552 and M. diernhoferi DSM 43524T at cell concentration of approximately 108, 107, 106, 105, 104, 103 and 102 CFU g-1 in white sand. (B) PCR-DGGE detection of M. gilvum VM0552 and M. diernhoferi DSM 43524T added in declining cell concentration together with a constant cell density of M. frederiksbergense LB501T of 108 CFU g-1. The symbol * indicates the detection limit on the figure.

- 73 - Chapter 2

Diversity analysis of fast-growing Mycobacterium populations in PAH contaminated soil. The MYCO-primer PCR-DGGE method was used to assess the Mycobacterium diversity in a set of PAH-contaminated soil samples with different contamination records (Table 2-2). Indigenous Mycobacterium cells could be detected in 6 of the 7 tested non-contaminated soils and in 6 of the 7 tested PAH-contaminated soils (Figure 2-3). Despite the high concentrations of PAHs, PCR-DGGE fingerprinting with universal eubacterial primers revealed, however, the presence of a heterogeneous bacterial soil community in soils negative in PCR with the MYCO-primer set. Moreover, DNA-extracts from parallel samples with added Mycobacterium cells produced good PCR products with the MYCO-primer set, omitting PCR inhibition as possible cause for the negative PCR results with the MYCO-primer set. The DGGE fingerprints of the Mycobacterium community in the positive soil samples were complex, comprising several bands for each soil (Figure 2-3). The 16S rRNA gene PCR products from 4 samples were randomly cloned and clones representing different bands from one soil fingerprint were sequenced. All sequences exhibited high levels of similarity to 16S rRNA gene sequences of Mycobacterium strains (Table 2-4) and could be placed within the phylogenetic group of fast-growing Mycobacterium species (Figure 2-4), confirming the specificity of the primer set. All positive soils revealed clone sequences most similar to the 16S rRNA genes of a variety of exclusively fast-growing Mycobacterium species. Sequences that were closely related to 16S rRNA gene sequences of known PAH- and oil-degrading species such as M. frederiksbergense, M. austroafricanum and M. petroleophilum were detected in soils K3840 and B101 but not in soil TM. However, the dominant number of 16S rRNA gene soil clone sequences from all 3 PAH-contaminated soils showed highest sequence similarity with the 16S rRNA gene of the relatively unknown M. tusciae. M. tusciae related sequences were not retrieved from the non- contaminated soil. Interestingly, the different soil fingerprints revealed bands closely related to M. tusciae but with different migration profiles. In addition, the cloned sequences showed a relatively high variation in similarity scores (from 99 to 95 %). The M. tusciae sequences isolated in this study grouped with other unidentified Mycobacterium sequences cloned from DNA from petroleum contaminated soils found by Cheung and Kinkle using the MycF and MycR primer pair (Cheung et al. 2001) (Figure 2-4). Besides the M. tuscia strains, only strains of the M. monacence

- 74 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

(AF107039) species, a fast-growing species represented by a type strain of clinical origin and an atypical isolate (U46146), seem to be closely linked to this cluster.

Low PAH con. High PAH con.

Mix S589 TB3 K3840 B101 E6068 TM

S589/4 S589/3 K3840/3 S589/6 S589/1 TM/8 K3840/4 TM/9 K3840/8 TM/2 M. frederiksb. K3840/2 TM/1 M. sphagni M. sp.WF2 K3840/1 B101/2 TM/3 M. gilvum K3840/6 B101/4 TM/6 S589/7 B101/7 TM/4 S589/5 B101/6 TM/5 B101/3 M. vaccae B101/5 TM/7 M. diernhoferi K3840/7 S589/2 B101/1

FIGURE 2-3 PCR-DGGE FINGERPRINT OF INDIGENOUS MYCOBACTERIUM CELLS IN SOIL SAMPLES USING MYCOBACTERIUM SPECIFIC PRIMERS MYCO66F AND GC40-MYCO600R Cloned ‘bands’ are indicated within the soil fingerprint based on the comparison of migration profiles of pure clones and the soil profile.

TABLE 2-4 CLONED 16S RRNA GENE SEQUENCES RETRIEVED FROM PAH-POLLUTED SOIL SAMPLES

NEAREST MATCH IN BLAST ANALYSIS ORIGIN CLONES ACCESSION N° SIMILARITY (ACCESSION N°) Soil S589 S589/1 / M. alvei DSM 44176T (AF023664) 97 % S589/2 / M. moriokaense DSM 44221T (AJ429044) 94 % S589/3 / M. lacus (AF406783) 97 % S589/4 / M. lacus (AF406783) 97 % S589/5 / M. anthracenicum * (Y15709) 98 % S589/6 / M. margeritense 1336 (AJ011335) 95 % S589/7 / M. hodleri DSM 44183T (X93184) 97 % Soil K3840 K3840/1 AY148216 Mycobacterium sp. M0183 (AF055332) 99 % K3840/2 AY148200 Mycobacterium sp. HXN1500 * (AJ457057) 98 % K3840/3 AY148207 M. tusciae DSM 44338T (AF058299) 95 % K3840/4 AY148201 M. frederiksbergense LB501T * (AJ245702) 98 % K3840/6 AY148214 M. austroafricanum DSM 44191T * (X93182) 98 % K3840/7 AY148197 M. gadium ATCC 27726 (X55594) 98 % K3840/8 AY148210 M. tusciae DSM 44338T (AF058299) 99 % Soil B101 B101/1 AY148202 Mycobacterium sp. JKD2385 (AF221088) 98 % B101/2 AY148208 M. isoniacini INA-I (X80768) 97 % B101/3 AY148198 M. holsaticum1406 (AJ310467) 97 % B101/4 AY148204 M. tusciae DSM 44338T (AF058299) 96 % B101/5 AY148212 M. septicum HX1900 (AJ457056) 99 % B101/6 AY148217 M. petroleophilum RF002 * (U90876) 98 % B101/7 AY148215 Mycobacterium. sp. WF2 * (U90877) 97 % Soil TM TM/1 AY148211 M. tusciae DSM 44338T (AF058299) 99 % TM/2 AY148209 M. tusciae DSM 44338T (AF058299) 99 % TM/3 AY148203 M. tusciae DSM 44338T (AF058299) 97 % TM/4 AY148205 M. tusciae DSM 44338T (AF058299) 97 % TM/5 AY148206 M. tusciae DSM 44338T (AF058299) 98 % TM/6 AY148196 M. moriokaense MCR07 (AF058299) 97 % TM/7 AY148213 M. septicum HX1900 (AJ457056) 98 % TM/8 AY148199 M. tusciae DSM 44338T (AF130308) 99 % * known oil or PAH degrading bacterium

- 75 - Chapter 2

17 16 15 14 13 12 11 10 9 8 7 6 5 4 3 2 1 0 X77779 R. gl ob eru lu s NC IMB12 3 15 100 X79290 Di. maris DSM43672 U90876 M. pe trol eo ph ilu m RF0 02 100 A Y14 82 17 M. sp . cl on e B1 01 /6 A J01 27 38 M. sp . D SM4 42 38 AF055332 M. sp. M0183 100 A Y14 82 16 M. sp . cl on e K3 84 0/1

77 A Y14 82 13 M. sp . cl on e TM/7 A J45 70 56 M. se pticu m HX N1 90 0 10 0 A Y14 82 12 M. sp . cl on e B1 01 /5 10 0 93 76 99 A J45 70 55 M. se pticu m HX N5 00 AF023664 M. al ve i DSM 44 17 6T AF221088 M. sp. JDK23855 92 100 100 A Y14 82 02 M. sp . cl on e B1 01 /1 96

75 X55591 M. kom oss ense DSM44078T 75 X55598 M. aichiense DSM44147T 100 A J01 26 26 M. sp. GP1 A J45 70 57 M. frederiksbergense HXN1500 A J27 62 74 M. fred eri ksb erg en se D SM4 43 46 T 85 AF294749 M. sp . cl on e KT-2 6 100 85 AF294750 M. sp . cl on e KT-2 7 A Y14 82 00 M. sp . cl on e K3 84 0/2 100 A Y14 82 01 M. sp . cl on e K3 84 0/4 8498

59 X55593 M. di ern ho feri D SM4 35 24 T M29564 M. ne oa ur um D SM44 07 4 T 85 A J31 04 67 M. ho lsa ticu m 14 06 100 A Y14 81 98 M. sp . cl on e B1 01 /3 99 83 X55594 M. ga di um ATC C2 77 26 10 0 96 A Y14 81 97 M. sp . cl on e K3 84 0/7 AF058712 M. pe re gri nu m D SM4 32 7 1T

82 X93184 M. ho dl eri DSM 44 18 3T 100 X80768 M. iso ni ac ini IN A-I 100 A Y14 82 08 M. sp . cl on e B1 01 /2 90 A Y14 81 96 M. sp . cl on e TM/6

91 X93033 M. mo rio kae nse MCR O 7

91 A B02 84 83 M. sp. TA5 100 AF330695 M. sp . cl on e C C-4 99 AF294742 M. sp . cl on e AC -1 X93183 M. parafortuitum DSM 43528T

77 X81996 M. gilvum DSM44503T X55595 M. au ru m D SM4 39 99 T 83 AF220431 M. sp . cl on e KT-1 9 100 87 X79094 M. ch lor op he no lic um D SM4 38 26 T A Y14 82 14 M. sp . cl on e K3 84 0/6 100 88 X93182 M. au stro afric an um D SM44 19 1T U90877 M. sp. WF2 90 A Y14 82 15 M. sp . cl on e B1 01 /7 A Y14 82 06 M. sp . cl on e TM/5

93 A Y14 82 04 M. sp . cl on e B1 01 /4 100 A Y14 82 05 M. sp . cl on e TM/4 93 96

89 A Y14 82 07 M. sp . cl on e K3 84 0/3 94 A Y14 82 03 M. sp . cl on e TM/3 94 AF058299 M. tusc iae DSM 44 33 8T

13 A Y14 82 11 M. sp . cl on e TM/1

7 A Y14 82 09 M. sp . cl on e TM/2 A Y14 81 99 M. sp . cl on e TM/8 100 94 A Y14 82 10 M. sp . cl on e K3 84 0/8 AF220428 M. sp . cl on e C T- 11

95 AF220430 M. sp . cl on e JT-1 5

96 AF107039 M. monacense B9-21-178 AF220427 M. sp . cl on e AT-3 97 100 U46146 M. sp. 98 92 AF220429 M. sp . cl on e C T- 12 100 AF220432 M. sp . cl on e KT-2 2 10 0 AF220433 M. sp . cl on e KT-2 3 AF294745 M. sp . cl on e C T- 24

66 AF294746 M. sp . cl on e C T- 25 AF294748 M. sp . cl on e JC -14 48 100 AF294744 M. sp . cl on e C T- 10 45 AF294747 M. sp . cl on e JC -13 100 AF294743 M. sp . cl on e C T- 9

- 76 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils.

FIGURE 2-4 PHYLOGENETIC ANALYSIS OF 16S RRNA GENE SEQUENCES DETECTED IN SOIL SAMPLES USING MYCOBACTERIUM SPECIFIC PRIMERS MYCO66F AND GC40-MYCO600R. Phylogenetic positioning of Mycobacterium 16S rRNA gene sequences detected in soil within the Mycobacterium genus. The evolutionary tree was generated by the Neighbor-joining method based on Kimara 2-parameter corrected similarity percentages of 538 bp 16S rRNA gene fragments between the MYCO primers and branching orders were evaluated using the Maximum-Parsimony algorithm. The topology was also evaluated by bootstrap analysis (500 reassemblings) and percentages of bootstrap support are indicated at the branch points, with values above 70% indicating reliable branches. An out- group of the closely related genera Rhodococcus and Dietzia was included to root the tree. The bar at the top indicates the estimated evolutionary distance, i.e., 1% indicating an average of 1 nucleotide substitution at any nucleotide position per 100 nucleotide positions. The evolutionary distance between two strains is the sum of the branch lengths between them.

DISCUSSION

We developed a specific PCR-DGGE method to rapidly assess the diversity of or monitor fast-growing Mycobacterium species in PAH-contaminated soil. From our test results, it could be concluded that the new MYCO66f and MYCO600r primer set, although theoretically also targeting a few slow-growing species, amplifies preferentially 16S rRNA gene of fast-growing Mycobacterium species from environmental samples. It is the first primer set that was specifically developed for the detection fast-growing Mycobacterium species only. All previously described primer combinations were specific for the total Mycobacterium genus (Böddinghaus et al. 1990; Kox et al. 1995; Kox et al. 1997; Talaat et al. 1997; Schwartz et al. 1998; van der Heijden et al. 1999; Mendum et al. 2000; Stinear et al. 2000; Cheung et al. 2001) or exclusively for pathogenic and facultative pathogenic slow-growing Mycobacterium species (De Beenhouwer et al. 1995; Schwartz et al. 1998). Even the Mycobacterium genus specific 16S rRNA gene primer set that was parallel designed during the course of this work and used in combination with Temperature Gradient Gel Electrophoresis (TGGE) for diversity analysis of indigenous Mycobacterium populations in petroleum-contaminated soil, was targeting both fast- and slow- growing species (Cheung et al. 2001). Forward primers MYCO66f (this study) and MycF (Cheung et al. 2001) were similarly conserved in 16S rRNA genes of Mycobacterium strains but reverse primer MYCO600r (this study) was far more specific for fast-growing Mycobacterium species than the MycR primer (Cheung et al. 2001).

- 77 - Chapter 2

DGGE analysis of gene fragments amplified with the MYCO-primer set could differentiate between most fast-growing Mycobacterium species including all important PAH-degrading species. For some very closely related species the DGGE fingerprints overlapped. For comparison, DGGE-differentiation of the same species was also limited when using PCR-products obtained with the Mycobacterium specific primer set of Cheung and Kinkle (Cheung et al. 2001). Similarly, TGGE-analysis of 16S rRNA gene fragments could not discriminate between several species of Burkolderia (Falcão Salles et al. 2002) and Bifidobacterium (Satokari et al. 2001) or other Gram-positive coryneform soil bacteria such as Arthrobacter and Nocardoides (Felske et al. 1999), due to the high levels of conservation of the amplified 16S rRNA gene fragments. It is clear that the practical resolution limit of the DGGE-technique is at the species or genus level or intermediate between the two, depending on the gene conservation level within the taxonomic group that is under investigation.

Most Mycobacterium strains were characterized by a single band DGGE-fingerprint, only a few showed satellite bands. The sequences of multiple bands for one strain displayed very high similarity. Others also reported multiple-band DGGE-patterns for pure strains of species such as Paenibacillus polymyxa (Nübel et al. 1996), Burkholderia cepacia (Falcão Salles et al. 2002) and Bifidobacterium adolescentis (Satokari et al. 2001) due to the presence of multiple 16S rRNA gene copies with sequence heterogeneity. Southern blotting and DNA-DNA hybridization revealed, however, that the indicated Mycobacterium strains contained a maximum of two copies of the rrn operon (Chapter 3), theoretically producing maximum 2 different bands on a DGGE-gel. This is consistent with other reports and 3 total genome sequences from Mycobacterium strains from clinical origin showing that slow- growing and fast-growing Mycobacterium strains possess only 1 or 2 copies of rrn genes respectively (Bercovier et al. 1986; Klappenbach et al. 2001). It is therefore unlikely that the obtained multiple band DGGE-fingerprints are due to the presence of more than 2 rRNA gene copies. Reaction conditions such as these used for the MYCO-primers, i.e., mixtures with high concentrations of very homologous 16S rRNA gene templates, may generate chimeras (Cole et al. 2003). However, it is unlikely that random chimera formation caused these multiple-band DGGE patterns since different DNA-preparations of the same strain always produced the same multiple-band DGGE-fingerprints. Moreover, cloned sequences of single strain

- 78 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils. multiple bands were negative in chimera analysis and showed limited variations. The slower migrating fainter satellite bands (located higher in the gel) may have been due to heteroduplex formation between 2 copies of 16S rRNA genes with 1 species. It is known that specific heteroduplexes can be detected in denaturing gels when 16S rRNA genes of Mycobacterium strains with at least 1 to 2 % difference in nucleotide sequence anneal (Waléria-Aleixo et al. 2000). In a mix of PCR products of 2 different genes, one or two heteroduplex bands additional to the one or two homoduplex bands may appear in the denaturing gel fingerprint. Alternatively, culture impurities could be the cause of multiple band patterns, although several culture DNA-extracts were tested and severe precautions for culture or template cross contamination were taken. Nevertheless, the possible appearance of multiple-band DGGE-fingerprints for single Mycobacterium strains may indicate that the number of different Mycobacterium species actually present in the soil community may well be lower than the number of bands in a soil fingerprint.

With the one-step PCR-DGGE method using only the MYCO66f and GC40- MYCO600r primer pair, we could detect Mycobacterium cells in white sand and several PAH-contaminated soils until a minimum cell concentration of circa 106 cells per gram soil. None of the other Mycobacterium genus specific primers sets developed in the past, report on the Mycobacterium abundance or detection limit in environmental samples for comparison (Schwartz et al. 1998; Mendum et al. 2000; Stinear et al. 2000; Cheung et al. 2001) but we found a similar detection limit when using the primer set developed by Cheung and Kinkle (Cheung et al. 2001). The value reported for a similar direct PCR-DGGE detection method for Burkholderia species in soil was only slightly lower (detection limit 5 × 105 CFU g-1) (Falcão Salles et al. 2002), although more copies of the rrn genes are present in the target bacterium (6 rrn copies in Burkholderia while 2 in Mycobacterium strains). Nested PCR using eubacterial primers in the first round and the MYCO-primers with GC-clamp in the second round could drastically lower the detection limit to circa 102 cells per gram soil. Another approach could be the use of the more abundant rRNA molecules instead of the rRNA gene as targets for the MYCO-primers in a reverse transcription PCR (RT-PCR) protocol. In a RT-PCR set up using the MYCO-primer set, Mycobacterium sp. LB501T was detected at a concentration as low as 102 active cells gr-1 soil (Hendrickx et al., unpublished data). Based on our results, all fast-growing

- 79 - Chapter 2

Mycobacterium strains are also expected to be detected equally well in a mixed Mycobacterium community. No differences in lysis or in preferential amplification based on primer homology would be expected between the different fast-growing PAH-degrading Mycobacterium species and there are indications that all fast-growing Mycobacterium species contain the same amount (2 copies) of 16S rRNA gene copies (see above).

Finally, the new specific PCR-DGGE method was successfully used to assess the diversity of fast-growing Mycobacterium species in a set of different contaminated and non-contaminated soils. Fast-growing Mycobacterium species could be detected in 6 out of the 7 tested PAH-contaminated soil samples and in 6 out of 7 tested non- contaminated soil samples. These results clearly suggest a wide distribution of fast- growing Mycobacterium species in the environment. For the 5 positive soils containing PAH-concentrations of 500 mg kg-1 or lower, there was no clear correlation between Mycobacterium biodiversity (assessed by the number of bands in the Mycobacterium DGGE fingerprints) and the PAH-concentration of the soils. Cheung and Kinkle (Cheung et al. 2001), however, observed a clear negative correlation between the number of Mycobacterium bands in the TGGE fingerprint (2 to 18) and the PAH-content (0.07 to 473 mg kg-1) of a soil. They suggested that the Mycobacterium diversity in more contaminated soil samples may be reduced by the toxicity of the PAHs or by a natural selection process in which the Mycobacterium community was enriched and finally dominated by one or a few populations more adapted to the higher PAH concentrations. The fact that there was no clear reduction in Mycobacterium diversity with increasing PAH-concentrations in our test soils could thus be due to the lack of toxicity or the lack of community adaptation in our soil samples with PAH-concentrations of 500 mg kg-1. With the one-step PCR-DGGE method, no Mycobacterium PCR signal could be detected in the 2 soils with highest PAH-concentrations of circa 1000 and 3000 mg kg-1. Although we did not perform toxicity tests, we have no indications that the very low or undetectable concentration of Mycobacterium species in heavily contaminated soils was due to toxicity as all tested soils were characterized by a rather complex total bacterial community DGGE- fingerprints obtained with universal primers and contained a total concentration of 106 to 108 cultivatable bacterial cells independent of the PAH-concentration (Vanbroekhoven et al. unpublished). The absence or lower concentrations of fast-

- 80 - Occurrence and diversity of Mycobacterium species in PAH-contaminated soils. growing Mycobacterium species in very heavily PAH-contaminated soils (1000 – 3000 mg kg-1) may indicate the natural selection of fast-growing Mycobacterium species in PAH-polluted soil enriched in poorly bioavailable and highly recalcitrant higher molecular PAHs. In soils containing high concentration of more easily degradable 3-ring PAHs Mycobacterium species might be out competed by more quickly growing PAH-degrading bacteria such as Sphingomonas or Pseudomonas species. Mycobacterium species are maybe better adapted to harsh oligotrophic soil conditions as they have a low maintenance energy demand and make use of several PAH bioavailability-enhancing mechanisms such as high-affinity uptake systems and adhesion to the substrate (Wick et al. 2001; Wick et al. 2002a).

Sequence analysis of the indigenous soil DGGE-fingerprints of 4 of the soils revealed the presence of strains closely related to known fast-growing PAH-degrading isolates from the M. frederiksbergense and M. austroafricanum species in 2 of the PAH- contaminated soils. None of the detected sequences seemed to originate from strains related to M. gilvum, another well-known PAH-degrading species (Boldrin et al. 1993; Bastiaens 1998; Bastiaens et al. 2000). However, sequences related to the M. tusciae species were repeatedly detected in all PAH-contaminated soils, originating from different countries and different industrial sites, but not in the non-contaminated soil. These results may indicate an important role for M. tusciae and/or related species in PAH-degradation processes in soil. The type strain of this species, M. tusciae strain DSM 44338T, is a facultative pathogenic clinical isolate from a sick child (Tortoli et al. 1999), but also 2 unpublished vinyl chloride degrading soil isolates (Coleman et al., unpublished) were recently identified as members of the M. tusciae species. Based on the different DGGE-bands and the varying similarity of the clones in our study to the type strain, we may have detected still unknown species relatively closely related to M. tusciae. Although, the M. tusciae species has never been isolated or detected before in PAH-contaminated soil, the M. tusciae sequences isolated in this study grouped with other unidentified Mycobacterium sequences cloned from DNA from petroleum contaminated soils found by Cheung and Kinkle using the MycF and MycR primer pair (Cheung et al. 2001).

- 81 - Chapter 2

The repeated detection of Mycobacterium cells in soils with low PAH-concentrations and low in organic carbon support the natural importance of fast-growing Mycobacterium species in PAH-polluted soil. The developed PCR-DGGE detection system is an important tool to specifically monitor the natural abundance, the diversity and the dynamics of these bacteria in soil for optimization of bioremediation. The developed primer pair may also be useful in a RT-PCR approach with ribosomal RNA soil extracts to analyze the diversity of the actively PAH-degrading population of fast- growing Mycobacterium species. Eventually these primers could be combined with primers developed for the detection of messenger RNA of the well conserved PAH catabolic genes in Mycobacterium strains (Khan et al. 2001; Krivobok et al. 2003). This will add to a better understanding of the role of Mycobacterium species in the biodegradation of PAHs in the environment.

ACKNOWLEDGEMENTS

This work was supported by the European Commission, through the contracts BIO4- CT97-2015 and QLRT-1999-00326. We thank E.M.H. Wellington for providing bacterial strains and S. Schioetz-Hansen, J. Amor and J. Vandenberghe for providing soil samples.

- 82 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

CHAPTER 3

MYCOBACTERIUM FREDERIKSBERGENSE, A MYCOBACTERIUM SPECIES SPECIALISED IN POLYCYCLIC AROMATIC HYDROCARBON (PAH) DEGRADATION, IS UBIQUITOUS IN PAH-CONTAMINATED SOILS

* REDRAFTED AFTER: LEYS NATALIE, RYNGAERT ANNEMIE, BASTIAENS LEEN, VAN CANNEYT MARK,

SWINGS JEAN, TOP EVA, VERSTRAETE WILLY, SPRINGAEL DIRK (SUBMITTTED) MYCOBACTERIUM

FREDERIKSBERGENSE, A MYCOBACTERIUM SPECIES SPECIALISED IN POLYCYCLIC AROMATIC

HYDROCARBON (PAH) DEGRADATION, IS UBIQUITOUS IN PAH-CONTAMINATED SOILS, ENVIRON.

MICROBIOL.

ABSTRACT

The fast-growing Mycobacterium species M. frederiksbergense seems to be specialized in colonizing PAH-contaminated soils and to play a role in PAH biodegradation in those soils. This species contained up to now only PAH-degrading isolates derived from PAH-contaminated soil. Based on 16S rRNA gene sequence similarity, FAME analysis and ribotyping, 7 additional PAH-degrading Mycobacterium isolates were identified as M. frederiksbergense strains. A culture- independent PCR based protocol was developed to detect and monitor specifically M. frederiksbergense strains in contaminated soils. A specific primer pair, MYCOFf and MYCOFr, was developed targeting exclusively the 16S rRNA gene sequence of M. frederiksbergense strains. PCR reactions using the new primer sets on Mycobacterium and non-Mycobacterium template DNA demonstrated that the MYCOFf and MYCOFr primer set was highly selective for M. frederiksbergense. Using the new primer set, M. frederiksbergense strains inoculated in soil could be detected at a cell concentration of 104 cells per g soil via direct PCR and subsequent DNA-DNA hybridization of the PCR products or at a cell concentration of maximum 102 cells per g soil via a nested PCR approach consisting of a first PCR reaction using universal primers and a second reaction using the MYCOFf/MYCOFr primer set. PCR on soil DNA extracts revealed the presence of M. frederiksbergens-like strains in PAH-

- 83 - Chapter 3 contaminated soils of diverse origin and different overall contamination profiles, PAH-concentrations and physico-chemical soil characteristics. These results indicate a role for M. frederiksbergense strains in the biological degradation of PAHs in PAH- contaminated environments.

INTRODUCTION

Polycyclic aromatic hydrocarbons (PAHs) are hazardous environmental pollutants that are found in high concentrations in the surface soil of sites housing former gas production plants and wood preservation plants through spills of coal tar and/or creosote (Cerniglia 1992). Biodegradation is considered as the main route of natural PAH removal in soil. Therefore, bioremediation can be used as an alternative for chemical or physical remediation techniques to clean up PAH contaminated sites. Many of the PAH-degrading bacteria isolated from PAH-contaminated soils have been identified as Mycobacterium (Bastiaens 1998; Bastiaens et al. 2000; Springael et al. unpublished). Those PAH-degrading Mycobacterium isolates were, based on 16S rRNA gene sequence, often assigned to the species M. frederiksbergense (Willumsen et al. 2001a), M. gilvum (Boldrin et al. 1993; Bastiaens et al. 2000; Vila et al. 2001; Gauthier et al. 2003), M. austroafricanum (Bogan et al. 2003), M. vanbaalenii (Heitkamp et al. 1988a; Godvidaswami et al. 1995; Wang et al. 1995; Khan et al. 2001; Moody et al. 2001; Khan et al. 2002), M. hodleri (Kleespies et al. 1996), M. flavescens (Dean-Ross et al. 1996), M. anthracenicum (Wang et al. unpublished) and M. chelonae (Kanaly et al. 2000a; Kanaly et al. 2000b; Kanaly et al. 2002). M. frederiksbergense is of particular interest for PAH-bioremediation purposes as up to now, this species seems to be specialized in PAH-degradation. The M. frederiksbergense species is taxonomically represented by only one strain, i.e., PAH- degrading type strain M. frederiksbergense FAn9T (DSM 44346T) (Willumsen et al. 2001a), but several PAH-degraders Mycobacterium isolates from diverse origin including Mycobacterium sp. strain LB501T, seem to be closely related to strain FAn9T based on 16S rRNA gene sequence (Bastiaens 1998; Bastiaens et al. 2000; Springael et al. unpublished). Strain LB501T displays several adaptations to the low bioavailability of PAH-compounds, i.e., it makes close contact with and strongly adhere to the PAH crystal surface, uses high-affinity uptake systems and has very low maintenance energy requirements (Wick et al. 2001; Wick et al. 2002a). Moreover, it

- 84 - Occurrence of M. frederiksbergense in PAH-contaminated soils. has a unique cell wall with an extreme high negative surface charge and high hydrophobicity, possibly playing a role in the modes of interaction of the strain with PAH-compounds (Bastiaens 1998; Bastiaens et al. 2000; Wick et al. 2001; Wick et al. 2002a; Wick et al. 2002b; Wick et al. 2003b). Furthermore, in contrast with other PAH-degrading Mycobacterium strains, several of those apparently FAn9T related strains are able to degrade PAHs at low temperature (e.g. 4-12°C) and in the presence of high salt concentrations (I = 0.500) (Leys et al. unpublished data). Direct detection of species such as M. frederiksbergense by PCR or other molecular techniques is therefore of importance for further study of the distribution and role of this species in PAH-contaminated environments. Primer sets exist that target all Mycobacterium species (Schwartz et al. 1998; Mendum et al. 2000; Cheung et al. 2001) or the group of the fast-growing Mycobacterium species (Chapter 2) for their detection by PCR in environmental samples. In 1996, 16S rRNA gene based primer set have been reported for the PCR detection of M. chlorphenolicum PCP1 (DSM 43826T) (Briglia et al. 1996) and PAH-degrading strains M. vanbaalenii PYR1 (DSM 7251T) and Mycobacterium sp. strain PAH135 (Wang et al. 1996). Specific detection methods for the PAH-degrading M. frederiksbergense FAn9T (DSM 44346T), however, do not exist. In this study, we report the development of a new species-specific detection method for selective monitoring of PAH-degrading strains of the M. frederiksbergense species. The new developed 16S rRNA gene primer set was used to assess the natural presence of M. frederiksbergense strains in a set of PAH-contaminated soils.

MATERIALS AND METHODS

Bacterial strains and growth conditions. The bacterial strains used in this study are described in Table 3-1. For DNA-extraction purposes, Mycobacterium strains were cultivated in Middelbrook 7H9 Broth (DIFCO) while all other strains were cultivated in 869-broth (Mergeay et al. 1985). All cultures were grown in the dark on an orbital horizontal shaker at 200 rpm at a constant temperature of 30 °C.

TABLE 3-1 STRAINS USED IN THIS STUDY

16S rRNA gene Strain Reference or origin MYCOF† accession N°

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MYCOBACTERIUM STRAINS M. aichiense 5545 T X55598 DSM 44147T - M. alvei CR-21 T AF023664 DSM 44176T - M. aurum 358 T X55595 DSM 43999T - M. austroafricanum E9789 T X93182 DSM 44191T - M. chlorophenolicum PCP-1 T X79094 DSM 43826T - M. diernhoferi SN1418 T X55593 DSM 43524T - M. frederiksbergense Fan9 T * AJ276274 DSM 44346T + M. gilvum BB1* X81891 DSM 9487 - M. gilvum SM35 T X81996 DSM 44503T - M. hodleri EM12 T * X93184 DSM 44183T - M. komossense Ko2 T X55591 DSM 44078T - M. neoaurum 3503 T M29564 DSM 44074T - M. parafortuitum 311 T X93183 DSM 43528T - M. peregrinum 6020 T AF058712 DSM 43271T - M. vanbaalenii PYR-1 T * U30662 DSM 7251T - Mycobacterium sp. GP1 AJ012626 (Poelarends et al. 1999) - Mycobacterium sp. HE5 AJ012738 DSM 44238 - Mycobacterium sp. LB208* AJ245704 (Bastiaens et al. 2000) - Mycobacterium sp. LB307T* AJ245703 (Bastiaens et al. 2000) - Mycobacterium sp. LB501T* AJ245702 (Bastiaens et al. 2000) + Mycobacterium sp. RF002* U90876 (Lloyd-Jones et al. unpublished) - Mycobacterium sp. VM0442* AF44636 (Springael et al. unpublished) - Mycobacterium sp. VM0447* AF44625 (Springael et al. unpublished) - Mycobacterium sp. VM0450* AF44623 (Springael et al. unpublished) - Mycobacterium sp. VM0451* AF44624 (Springael et al. unpublished) - Mycobacterium sp. VM0452* AF44626 (Springael et al. unpublished) - Mycobacterium sp. VM0456* AF44622 (Springael et al. unpublished) - Mycobacterium sp. VM0458* AF44630 (Springael et al. unpublished) + Mycobacterium sp. VM0503* AF44628 (Springael et al. unpublished) + Mycobacterium sp. VM0504* AF44634 (Springael et al. unpublished) - Mycobacterium sp. VM0505* AF44633 (Springael et al. unpublished) - Mycobacterium sp. VM0531* AF44629 (Springael et al. unpublished) + Mycobacterium sp. VM0552* AF44635 (Springael et al. unpublished) - Mycobacterium sp. VM0573* AF44627 (Springael et al. unpublished) - Mycobacterium sp. VM0579* AF44631 (Springael et al. unpublished) + Mycobacterium sp. VM0583* AF44637 (Springael et al. unpublished) - Mycobacterium sp. VM0585* AF44632 (Springael et al. unpublished) + Mycobacterium sp. VM0587* AF44638 (Springael et al. unpublished) - Mycobacterium sp. VM0588* AF44639 (Springael et al. unpublished) - Mycobacterium sp. WF2* U90877 (Lloyd-Jones et al. unpublished) - OTHER GENERA - Rhodococcus erythropolis ICPB 4417 T X81929 DSM 43066T - Nocardia asteroides N3 NR (Springael et al. unpublished) - Dietzia maris VM0283 NR (Wattiau et al. 1999) - Dietzia maris IMV 195 T X79290 DSM 43627T - Actinosynnema mirum 101 T X84447 DSM 43827T - Arthrobacter sulfureus 8-3 T X83409 DSM 20167T - Planomonospora parontospora B-987 T AB028653 DSM 43869T - Promicromonospora citrea INMI 18 T X83809 DSM 43110T - Streptomyces aureofaciens A-377 T NR DSM 40127T - Streptomyces rutgersensis BJ-608 T NR DSM 40830T - Streptomyces phaeofaciens T-23 T D44381 DSM 40367T - Brevundimonas diminuta 342 T AJ227778 DSM 7234T - Sphingomonas chlorophenolicum T X87161 DSM 7098T - Pseudomonas putida NR DSM 8368 - * PAH degrading strains † result of PCR with primers MYCOFf and MYCOFr: + = detectable PCR product, - = no detectable PCR product.

Soils used in this study. The soil samples used in this study were taken from different anthropogenic PAH-contaminated sites (Table 3-2). Chemical properties of the soil samples were analyzed as described previously (Chapter 2).

TABLE 3-2 SOIL SAMPLES USED IN THIS STUDY

- 86 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

Soil TOC PAH conc. Oil conc. DNA conc.* MYCO† MYCOF†† Soil Origin pH type (%) (mg kg-1) (mg kg-1) (µg g-1) PCR, cells g-1 PCR, cells g-1 K3840 Gasoline station site (Denmark) Sand 8.20 0.50 20 98 2.75 + , 109 + , 109 B101 Coal gasification plant (Belgium) Sand 7.00 2.63 107 70 27.25 + , 108 + , 108 E6068 Gasoline station site (Denmark) Sand 7.96 9.94 258 300 5.40 +, 107 NP, <106 TM Coal gasification plant (Belgium) Sand 8.00 3.85 506 4600 4.75 + , 108 + , 2x106 Barl Coal gasification plant (Germany) Gravel 8.90 4.63 1029 109 6.15 NP NP, <106 AndE Railway station site (Spain) Clay 8.10 2.35 3022 2700 3.40 +, <106 NP, <106 * DNA recovery per g soil, mean value of 2 parallel extractions of 1 soil sample. † result of nested PCR on soil DNA extract combining eubacterial primers 27f and 1492r with Mycobacterium specific primers MYC066f and MYCO600r (Chapter 2): + = PCR product, NP = no detectable PCR product in agarose electrophoresis of PCR products †† result of direct PCR on soil DNA extract using M. frederiksbergense specific primers MYCOFf and MYCOFr: + = PCR product, NP = no detectable PCR product in agarose electrophoresis of PCR products; cells g-1 = estimated Mycobacterium cell content based on a direct MPN-PCR approach using only specific primers.

DNA-extraction. Extraction of genomic DNA from pure bacterial cultures and soil samples was performed as described previously (Chapter 2). Phylogenetic analysis of M. frederiksbergense related Mycobacterium strains. A multiple alignment of Mycobacterium 16s rRNA gene sequences was constructed using the Bionumerics software (Version 2.50, Applied Maths, Belgium). The alignment consisted of approximately 200 sequences of both fast- and slow growing Mycobacterium species available from the GenBank database (NCBI) with a continuous stretch of circa 1300 bp. Sequences were edited manually to remove nucleotide positions of ambiguous alignment and gaps. Sequence similarities were calculated over the total length of the 16S rRNA gene and corrected using Kimura's two-parameter algorithm to compensate for multiple nucleotide exchange and a distance-based evolutionary tree was constructed using Kimura's corrected similarity values in the Neighbor-Joining algorithm (Saitou et al. 1987). The topography of the branching order within the dendrogram was evaluated by using the Maximum- Likelihood and the Maximum-Parsimony character-based algorithms in parallel combined with bootstrap analysis with a round of 500 reassemblings. An out-group of the closely related genera Rhodococcus and Dietzia was included to root the tree. Ribotyping of Mycobacterium strains. Ribotyping was performed, to identify the number and arrangement of 16S rRNA gene copies in the chromosome, a species or strain specific fingerprint. Chromosomal DNA from pure bacterial cultures was digested with restriction enzyme EcoRV (GibcoBRL) and the DNA fragments were separated by electrophoresis in 1% agarose. Fragments were transferred to Hybond-N+ membrane (Amersham Biosciences Europe GmbH Benelux, Roosendaal, The Netherlands) by Southern blotting, and hybridizations were carried out according to established protocols (Sambrook et al. 1989). The 538bp amplicons produced with the MYCO66f and MYCO600r primer set (Chapter 2) from Mycobacterium sp. strains LB307T and LB501T (Table 3-1) were chemiluminescently labeled using dioxigenin (DIG) (Boehringer Mannheim - Roche Diagnostics, Vilvoorde, Belgium) according to the supplier’s protocols, and subsequently simultaneously used as probes. Most strains were ribotyped at least twice to ensure the reproducibility of the fingerprint patterns. Hybridization patterns were compared for pattern similarity using the Bionumerics software (Version 2.50, Applied Maths, Belgium). Number of theoretical cutting sites was calculated using the REMAP program of the EMBOSS software.

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FAME analysis. Whole-cell fatty acid analysis (FAME) was performed on cells grown at 28°C for 24h on Tryptone Soya Agar (TSA-FAA) medium. Cellular fatty acids were saponified, methylated, extracted and analyzed by gas chromatography following the procedures given for the Sherlock Microbial Identification System (MIDI, Inc., Delaware U.S.A.). Identifications and comparisons were made by using the MIDI Aerobe database (TSBA version 3.10). Design of a M. frederiksbergense group specific 16S rRNA gene primer set. Specific primers were selected in highly variable regions from the constructed alignment of Mycobacterium 16S rRNA genes. The forward primer MYCOFf (5’ CCGAATATGACCATGCACTTCC 3’, E. coli numbering bp 179- 201) and reverse primer MYCOFr (5’ AAGGGAAACCACATCTCTGCAGT 3’, E. coli numbering bp 996-1019) were selected to amplify specifically an 840 bp fragment of the 16S rRNA gene of M. frederiksbergense and close relatives. Candidate primers were analyzed with Blast (Genebank, NCBI) (Altschul et al. 1990) and Sequence Match (RDPII) (Cole et al. 2003) software to select the best and most selective primer combination with no or limited mismatches within the group of M. frederiksbergense related strains and low homology with other strains. PCR amplification of 16S rRNA genes from pure strain and soil DNA. To assure that the soil DNA was of good quality for PCR, dilution series of all soil DNA extracts were tested in PCR with eubacterial 16S rRNA gene primer pair GC40-63f and 518r as described previously (Marchesi et al. 1998). For nested PCR protocols eubacterial primers 27fC (5’ AGAGTTTGATCCTGGCTCAG 3’) and 1492rC (5’ TACGGCTACCTTGTTTACGACTT 3’) were used in the first amplification round as described elsewhere (Polz et al. 1998). The PCR protocol used with the MYCOFf and MYCOFr primer pair consisted of an initial short denaturation of 15 s at 95°C, followed by 50 cycles of one denaturation step for 3 s at 95°C, one annealing step for 10 s at 65°C and one elongation step for 30 s at 74°C. The last step included an extension for 2 min at 74°C. PCR was performed on Biometra or Perking Elmer PCR-machines. PCR mixtures contained 100 ng of pure strain DNA, 1µl of dilutions of soil DNA or 1µl of first-round PCR product as templates, 1 U Taq polymerase, 25 pmol of the forward primer, 25 pmol of the reverse primer, 10 nmol of each dNTP, and 1 × PCR buffer in a final volume of 50 µl. Primers were synthesized by Westburg (Westburg BV, Leusden, The Netherlands) and the Taq polymerase, dNTPs and PCR buffer were purchased from TaKaRa (TaKaRa Ex TaqTM, TaKaRa Shuzo Co., Japan). Agarose gel electrophoresis of PCR products was run in 1.5% agarose gels in 1x EY buffer with 50µg l-1 ethidiumbromide for 1 hour at 90V. Sensitivity of PCR detection method. To study the sensitivity of the PCR method, a known amount of viable M. frederiksbergense LB501T cells were added to soil at different final cell concentrations (i.e. a 10-fold dilution series of approximately 108 to 101 cells g-1) prior to DNA-extraction. Inoculum cells were harvested from liquid cultures, washed twice and added in 100 µl aqueous suspensions to 1 g of soil. The total soil DNA was subsequently used as template for PCR with the MYCOF-primers. PCR products were analyzed by electrophoresis in a 1.5% agarose gel (MetaPhor, BioWhittaker, Labtrade Inc., Miami, Florida, USA), transferred to a Hybond-N+ membrane (Amersham Biosciences Europe GmbH Benelux, Roosendaal, The Netherlands) by Southern blotting, and hybridized with dioxigenin chemiluminescent labeled amplicons (Boehringer Mannheim, Roche Diagnostics Vilvoorde, Belgium) produced with the MYCOF primer set from strain LB501T, according to established protocols

- 88 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

(Sambrook et al. 1989). Detection of DIG probes was performed with a DIG luminescence detection kit (Boehringer Mannheim, Roche Diagnostics Vilvoorde, Belgium) following the suppliers protocol. Estimation of cell concentration. To estimate the cell concentration naturally present in soil samples a ‘dilution to extension PCR’ approach was used, i.e., soil DNA extracts were diluted 1:1, 1:10, 1:100 and 1:1000 times and used as template in PCR. The final cell concentration within a soil was deduced from the highest template dilution for which a PCR product was still detected, taking into account that the highest dilution giving a signal contained a cell density approaching the determined detection limit. Parallel soil samples with added cells were regarded as positive PCR controls to assure that negative PCR results with uninoculated samples were not due to PCR inhibition effects. Sequence analysis of amplified 16S rRNA gene fragments. PCR products obtained with primer set MYCOFf and MYCOFr were cloned into plasmid vector pCR2.1-TOPO using the TOPO Cloning Kit (NV Invitrogen SA, Merelbeke, Belgium) as described by the manufacturer. For a selection of the clones, a 500 bp long fragment of the inserts was sequenced (Westburg BV, Leusden, The Netherlands). A similarity analysis of the 16S rRNA gene sequences was done using the Advanced Blast Search program available from Genbank (NCBI) (Altschul et al. 1990). Nucleotide 16S RNA gene sequence accession numbers. The partial DNA sequence of Mycobacterium 16S RNA gene clones reported in this study are available from Genbank under accession numbers AY345102 to AY345119.

RESULTS

Phylogenetic analysis of M. frederiksbergense related PAH-degrading strains. The new constructed phylogentic tree confirmed the close relationship between the species M. frederiksbergense and the species M. diernhoferi, M. neoaurum and PAH- degrading M. hodleri (Figure 3-1, bottom part of tree), as was previously shown (Willumsen et al. 2001a). Based on 16S rRNA gene sequence similarity, the PAH- degrading type strain FAn9T of the M. frederiksbergense species (described in 2001) is the closest relative to 7 other PAH-degrading Mycobacterium isolates isolated from freshwater sediment (strain CH-1) or soil (strains LB501T, VM503, VM458, VM531, VM585, and VM579) (Figure 3-1). The next most related branch consists of only 3 unidentified isolates from a deep-sea sediment (strain JoyU) and from air biofilters capable of degrading toluene (strain LAB2) or linear n-alkanes (strain HXN1500). The 16S rRNA-genes of the M. frederiksbergense related strains are similar but still significant different from each other, unlike M. gilvum related PAH-degrading strains which show identical 16S rRNA genes.

- 89 - Chapter 3 4 3 2 1 0 X79290 Dietzia maris DSM43672 X52923 Mycobacterium gordonae ATCC14470T AF480589 Mycobacterium poriferae ATCC35087T 98 U90877 Mycobacterium sp. WF2 * 95 X55596 Mycobacterium chubuense ATCC27278T 100 97 X79094 Mycobacterium chlorophenolicum DSM43826T

99 AF44638 Mycobacterium sp. VM0587 * 100 AF44639 Mycobacterium sp. VM0588 * AF44623 Mycobacterium sp. VM0450 * 97 100 100 U30662 Mycobacterium vanbaalenii DSM7251T * AF44624 Mycobacterium sp. VM0451 * 98 AF44622 Mycobacterium sp. VM0456 * 100 AF190800 Mycobacterium austroafricanum CIP-I-2126 101000 100 X93182 Mycobacterium austroafricanum DSM44191T

100 AF44625 Mycobacterium sp. VM0447 *

97 100 AF44626 Mycobacterium sp. VM0452 * AF44627 Mycobacterium sp. VM0573 * U90876 Mycobacterium sp. RF002 *

100 AJ012738 Mycobacterium sp. DSM44238

100 AJ245704 Mycobacterium sp. LB208 *

0 X81996 Mycobacterium gilvum DSM44503T

0 X81891 Mycobacterium gilvum DSM9487 AJ245703 Mycobacterium sp. LB307T * 0 100 AF44636 Mycobacterium sp. VM0442 * 98 0 AF44633 Mycobacterium sp. VM0505 * 0 AF44634 Mycobacterium sp. VM0504 * 0 AF44635 Mycobacterium sp. VM0552 * 100 AF44637 Mycobacterium sp. VM0583 * X93183 Mycobacterium parafortuitum DSM43528T X55598 Mycobacterium aichiense DSM44147T 96 AF023664 Mycobacterium alvei DSM44176T 96 X55591 Mycobacterium komossense DSM44078T 100 AF058299 Mycobacterium tusciae DSM44338T 96 AF058712 Mycobacterium peregrinum DSM43271T X93184 Mycobacterium hodleri DSM44183T 98 AF268445 Mycobacterium neoaurum 98 100 AF480582 Mycobacterium 'lacticola' ATCC9626

98 AF284430 Mycobacterium sp. ESD X55593 Mycobacterium diernhoferi DSM43524T 98 AJ007009 Mycobacterium sp. LAB2

100 98 AB010910 Mycobacterium sp. JoyU 100 AJ457057 Mycobacterium sp. HXN1500 AJ276274 Mycobacterium frederiksbergense DSM44346T * 98 AF44628 Mycobacterium sp. VM0503 * AF054278 Mycobacterium sp. CH-1 * 100 93 AJ245702 Mycobacterium sp. LB501T * AF44630 Mycobacterium sp. VM0458 * 96 AF44629 Mycobacterium sp. VM0531 * 99 AF44632 Mycobacterium sp. VM0585 * 100 AF44631 Mycobacterium sp. VM0579 *

FIGURE 3-1 PHYLOGENETIC ANALYSIS OF M. FREDERIKSBERGENSE-RELATED PAH- DEGRADING MYCOBACTERIUM STRAINS BASED ON 16S RRNA GENE SEQUENCE An evolutionary tree of the total Mycobacterium genus was generated by the Neighbor-joining method based on Kimara 2-parameter corrected similarity percentages and branching orders were evaluated using the Maximum-Parsimony algorithm. The topology was also evaluated by bootstrap analysis (500 reassemblings) and percentages of bootstrap support are indicated at the branch points, with values above 70% indicating reliable branches. An out-group of the closely related genera Rhodococcus and Dietzia was included to root the tree. The bar at the top indicates the estimated evolutionary distance, i.e., 1% indicating an average of 1 nucleotide substitution at any nucleotide position per 100 nucleotide positions. The evolutionary distance between two strains is the sum of the branch lengths between them. All branches in the Mycobacterium genus tree were collapsed except the branch of the M. frederiksbergense species. Isolates capable of degrading PAHs are indicated with an *. The cluster of proposed M. frederiksbergense strains is highlighted in a box.

- 90 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

The 8 M. frederiksbergense related strains showed similar ribotyping band patterns, which were clearly different from those of other Mycobacterium species, supporting their close phylogenetic relationship (Figure 3-2). Theoretically EcoRV was expected to cut once in the 16S rRNA gene of FAn9, LB501T and VM503 and twice in the 16S rRNA gene of VM531, VM458, VM579 and VM585. Therefore, the observed 3 to 4 bands in the patterns for FAn9, LB501T and VM503 and 4 to 6 bands in the patterns for VM531, VM458, VM579 and VM585 (Figure 3-2) would indicate at least two different copies of the 16S rRNA gene. FAME analysis further confirmed the grouping of the 8 PAH-degrading Mycobacterium strains in one cluster similar to the cluster based on 16S rRNA gene sequence similarity (data not shown). Thus, 16S rRNA gene similarity, ribotyping and FAME analysis strongly suggest that strains LB501T, CH-1, VM503, VM531, VM458, VM579 and VM585 should be assigned to the M. frederiksbergense species.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

M. frederiksbergense M. gilvum M. austroafricanum related strains related strains related strains

T FIGURE 3-2 RIBOTYPING PATTERNS OF M. FREDERIKSBERGENSE FAN9 RELATED PAH- DEGRADING MYCOBACTERIUM STRAINS AND OTHER MYCOBACTERIUM SPECIES Lanes: 1, M. frederiksbergense FAn9T; 2, Mycobacterium sp LB501T; 3, Mycobacterium sp. VM531; 4, Mycobacterium sp. VM585; 5, Mycobacterium sp. VM458; 6, Mycobacterium sp. VM579; 7, Mycobacterium sp. VM503; 8, Mycobacterium sp. LB307T; 9, Mycobacterium sp.VM552; 10, Mycobacterium sp. VM505; 11, Mycobacterium sp. VM583; 12, Mycobacterium sp. VM504; 13, Mycobacterium sp. VM442; 14, M. vanbaalenii PYR1T; 15, Mycobacterium sp.VM451; 16, Mycobacterium sp. VM456; 17, Mycobacterium sp. VM447; 18, Mycobacterium sp. VM573; 19, , Mycobacterium sp. VM450. For the tested strains the ribotyping patterns were divided in 3 groups supporting the phylogenetic division based on 16S rRNA gene similarities.

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Species specific detection of PAH-degrading M. frederiksbergense strains. Primer set MYCOFf and MYCOFr was selected to selectively amplify an 840 bp fragment of the 16S rRNA gene of M. frederiksbergense related strains only. The forward primer MYCOFf was 100% conserved in only 6 out of the circa 900 Mycobacterium 16S rRNA gene sequences currently available in the NCBI database, i.e., the sequences of M. frederiksbergense FAn9T, and related strains LB501T and VM458, but also of HX1500, JoyU and M. alvei (Figure 3-3). Two mismatches were found in the 16S rRNA genes of the other related strains VM0503, VM0531 and CH- 1 (Figure 3-3). Due to unidentified bases in the corresponding primer region of the 16S rRNA gene, primer-sequence mismatches found for strains LAB2 and VM0585, VM0579 could not be evaluated (Figure 3-3). Reverse primer MYCOFr was 100% similar in DNA sequence to 40 16S rRNA gene sequences, of which 19 were from Mycobacterium strains, 19 from Rhodococcus strains and 2 from Bifidobacterium strains. Homologous primer regions were found in 16S rRNA gene sequences of M. frederiksbergense strains LB501T, VM503, CH-1, VM458, VM585, VM531 and VM579 (Figure 3-3). In addition, the reverse primer was also conserved in several M. fortuitum, M. celatum, M. flavescens, M. novocastrense, M. poriferae, M. brumae, M. acapulcensis and different unclassified Mycobacterium sp. strains. Two mismatches were found in the primer region of M. frederiksbergense FAn9T and 6 mismatches for strains Mycobacterium sp. LAB2 and JoyU (Figure 3-3). In PCR, primers set MYCOFf and MYCOFr proved to be M. frederiksbergense species specific. Of the 35 tested Mycobacterium strains representing 14 different species, only 7 reacted positively and produced PCR products of the expected size, i.e., strains FAn9T, LB501T, VM503, VM585, VM458, VM579, and VM531. Strains LAB2, JoyU and HXN1500 were not available and thus not tested, but we predicted that 16S rRNA genes of these strains would not react with the MYCOF-primer pair based on the high number of mismatches with the reverse primer. The DNA from strains belonging to the closest neighbor cluster such as M. diernhoferi and M. neoaurum did not amplify with the primers. Negative PCR-results were also obtained in PCR-reactions with template DNA from all other bacteria tested (Table 3-1).

- 92 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

MYCOFf MYCOFr 5’.CCGAATATGACCATGCACTTCC 3’ 3’.TGACGTCTCTACACCAAAGGGAA…5’

M. fredericksbergense (AJ276274) ------C------G------Mycobacterium sp. LB501T (AJ245702) ------Mycobacterium sp. CH1 (AF054278) ------G-C------Mycobacterium sp. VM0503 (AF44628) ------G-C------Mycobacterium sp. VM0458 (AF44630) ------MYK-Y------R- SRK------Mycobacterium sp. VM0585 (AF44632) ------R-Y------MYK-Y------R- SRK------Mycobacterium sp. VM0579 (AF44631) ------R-Y------MYK-Y------R- SRK------Mycobacterium sp. VM0531 (AF44629) ------G-C------MYK-Y------R- SRK------Mycobacterium sp. LAB2 (AJ007009) ------M------G--T-A------T- A- C------Mycobacterium sp. JoyU (AB010910) ------G--T-A------T- A- C------Mycobacterium sp. HXN1500 (AJ457057) ------G--T-A------T- A- C------M. diernhoferi (X55593) ------GC----T------A GAC-A------A-GTC------M. neoaurum (M29564) ------T------CGG------A GAC-A------A-GTC------M. aurum (X55595) ------G------T---C------T. G-A ACGGT------TC-AT------M. vanvaalenii (U30662) ------CAC-----T--CTGGC-G- ACGGT------TC-AT------M. austroafricanum (X93182) ------CAC-----T--CTGGC-G- ACGGT------TC-AT------M. chlorophenolicum (X79094) ------G------C----GT-G-A G--C------GCC------M. gilvum (X81996) ------G------GCATG------A G--C------GCC------

FIGURE 3-3 DNA-SEQUENCE HOMOLOGY BETWEEN THE MYCOF PRIMER SET AND 16S RRNA GENE SEQUENCES FROM DIFFERENT MYCOBACTERIUM SPECIES Results are presented in a consensus table of matches. Dashes indicate identical nucleotides.

Detection limit for PCR-based detection of M. frederiksbergense. In a direct PCR protocol, the primer set MYCOFf and MYCOFr, allowed the detection of inoculated Mycobacterium sp. strain LB501T in PAH-contaminated soils at a cell density of 106 cells g-1 of soil. Addition of formamide, ureum, glycerol, DMSO or PEG in the PCR reaction, could not improve the detection limit. The detection limit could be decreased to at least 104 cells g-1 of soil by means of Southern blot hybridization of the PCR products with the 16S rRNA gene PCR product obtained from LB501T genomic DNA as a probe. By combining the MYCOF primer set with eubacterial primers 27f and 1492r in a nested PCR protocol, the detection limit was lowered to maximum 102 cells per g of soil. The nested PCR approach was preferred as it was the the least laborious and most sensitive.

Screening PAH-contaminated soils for the presence of M. frederiksbergense strains. The MYCOF primer set was used to screen different contaminated soil samples with different contamination records (Table 3-2) for the presence of M. frederiksbergense related strains. The primer set detected indigenous M. frederiksbergense related cells in 3 out of the 6 tested PAH-contaminated soils. Sandy soils K3840, B101 and TM polluted with relatively low concentrations of PAHs tested positively, while sandy

- 93 - Chapter 3 soil E6068 and highly contaminated gravel soil Barl and clay soil AndE were negative. Sequencing of the obtained 16S rRNA gene products retrieved from soil TM (Table 3-2), confirmed that the detected signals corresponded indeed to M. frederiksbergense related strains. All 9 randomly selected clone sequences were 99 tot 100 % similar with the 16S rRNA gene sequences of M frederiksbergense type strain Fan9T or the most closely related strains, LB501T or CH-1 (Table 3-3). The estimated concentration of M. frederiksbergense cells in the positive soils was relatively high, i.e., circa 106-109 cells per gram of soil (Table 3-2).

TABLE 3-3 CLONED SEQUENCES RETRIEVED FROM PAH-POLLUTED SOIL SAMPLES

ORIGIN CLONES ACCESSION N° NEAREST MATCH IN BLAST ANALYSIS (ACCESSION N°) Soil TM TM/LB1 AY345102 100% M. frederiksbergense DSM 44346T (AJ276274) TM/LB2 AY345103 99% M. sp. CH-1 (AF054278) TM/LB3 AY345104 100% M. frederiksbergense DSM 44346T (AJ276274) TM/LB4 AY345105 100% M. frederiksbergense DSM 44346T (AJ276274) TM/LB5 AY345106 100% M. frederiksbergense DSM 44346T (AJ276274) TM/LB6 AY345107 100% M. frederiksbergense DSM 44346T (AJ276274) TM/LB7 AY345108 99% M. sp. CH-1 (AF054278) TM/LB8 AY345109 99% M. frederiksbergense DSM 44346T (AJ276274) TM/LB9 AY345110 100% M. frederiksbergense DSM 44346T (AJ276274) * known oil or PAH degrading bacterium

DISCUSSION

The M. frederiksbergense species is a relatively recently described Mycobacterium species with FAn9T (DSM 44346T) as type strain. FAn9T was isolated from a Danish PAH-contaminated soil and able to mineralize phenanthrene, fluoranthene and pyrene (Willumsen et al. 2001a). As more PAH-degrading Mycobacterium strains have been isolated, the M. frederiksbergense species is becoming more and more interesting. A phylogenetic analysis showed that so far all described close relatives of the species types strain seem to have the capacity to degrade PAHs. In this paper, the close relationship of strains in the M. frederiksbergse branch of the 16S rRNA gene based phylogenetical tree was confirmed by FAME analysis and ribotyping, so that seven other PAH-degrading isolates could be assigned to the M. frederiksbergse species with almost 100% certainty. The M. frederiksbergense related PAH-degrading strains were shown to contain very similar but not identical 16S rRNA genes. DNA-DNA hybridization should be performed to further confirm this.

- 94 - Occurrence of M. frederiksbergense in PAH-contaminated soils.

A specific detection method to rapidly assess the presence and behavior of M. frederiksbergense strains was non-existing but is essential for future use in bioremediation projects. The new primer set MYCOFf and MYCOFr could selectively detect M. frederiksbergense-related Mycobacterium strains in soil samples to at least 102 cells per gram of soil in a nested PCR approach. Indigenous M. frederiksbergense related cells were detected in 3 out of the 6 PAH-contaminated soils tested. Using a primer set targeting the 16S rRNA gene of all fast-growing Mycobacterium species, it was shown that all tested soils were colonized with relatively high numbers of fast-growing Mycobacterium cells (108–109 cells g-1 soil) and a quite heterogeneous Mycobacterium population (Chapter 2). Using this genus specific primer set, however, M. frederiksbergense related sequences were only retrieved from soil K3840 (Chapter 2). In this study, highest concentrations of M. frederiksbergense strains (109 cells g-1) were again detected in soil K3840 but also in 2 other soils. Detection methods targeting a larger group of Mycobacterium species are biased in favor of the most abundant populations, which can completely mask the detection of other less abundant species (Chapter 2). A strain specific PCR-based detection method is thus really valuable and can reveal additional information on the Mycobacterium population detecting species that would otherwise be missed. The relatively high cell concentrations and the frequent occurrence of M. frederiksbergense related Mycobacterium strains in some PAH-polluted soils clearly suggest an important role of these bacteria in the natural biorestauration of contaminated sites. Soils containing M. frederiksbergense related cells were characterized with relatively low concentrations of PAHs (< 100 mg kg-1) or high concentrations of oil (4600 mg kg-1). The detection of higher concentrations of M. frederiksbergense cells in PAH-polluted soils containing lower concentrations of PAHs may confirm our hypothesis of natural selection of fast-growing Mycobacterium species in soils enriched in poorly bioavailable and highly recalcitrant higher molecular PAHs (Chapter 2). In soils containing high concentrations of more easily available and degradable 3-ring PAHs (mainly phenanthrene), Mycobacterium strains might loose the competition with faster growing PAH-degrading Sphingomonas strains which have been detected in high concentrations in these soils (Chapter 4). On the other hand, the more slowly growing Mycobacterium strains may win the competition in more harsh environmental conditions with low carbon and other nutrient concentrations prevailing in soils in a later stadium of biorestauration

- 95 - Chapter 3 containing lower PAH-concentrations of mainly higher molecular PAHs. Mycobacterium strains, and especially the M. frederiksbergense strains, have shown to be specialized in degradation of higher molecular PAHs, to be able to shift to low maintenance energy and to make use of several PAH-bioavailability-enhancing mechanisms such as high-affinity uptake systems and adhesion to the substrate (Wick et al. 2001; Wick et al. 2002a).

In conclusion, the M. frederiksbergense species specific PCR detection method described in this study is indispensable to further study the distribution and contribution of this specific group of Mycobacterium strains in the PAH removal in environmental samples. The newly developed 16S rRNA gene primers may also be useful in a RT-PCR approach targeting the rRNA in order to identify the active strains involved in PAH biodegradation in the environment.

- 96 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

CHAPTER 4

OCCURRENCE AND PHYLOGENETIC DIVERSITY OF SPHINGOMONAS IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS)

* REDRAFTED AFTER: LEYS NATALIE, RYNGAERT ANNEMIE, BASTIAENS LEEN, TOP EVA, VERSTRAETE

WILLY, SPRINGAEL DIRK (2004) OCCURRENCE AND PHYLOGENETIC DIVERSITY OF SPHINGOMONAS IN

SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS), APPL. ENVIRON.

MICROBIOL. 70 (4): 1944– 955.

ABSTRACT

Bacterial strains of the Sphingomonas genus are often isolated from contaminated soils for their ability to use polycyclic aromatic hydrocarbons (PAH) as sole source of carbon and energy. The direct detection of Sphingomonas strains in contaminated soils, either indigenous or inoculated, is as such of interest for bioremediation purposes. In this study, a culture-independent PCR based detection method using specific primers targeting the Sphingomonas 16S rRNA gene, combined with Denaturing Gradient Gel Electrophoresis (DGGE), was developed to asses their diversity in PAHs contaminated soils. PCR using the new primer pair on a set of template DNAs of different bacterial genera showed that the method was selective for bacteria belonging to the Sphingomonadaceae family. Single-band DGGE profiles were obtained for most Sphingomonas strains tested. Strains belonging to the same species had identical DGGE fingerprints and in most cases these fingerprints were typical for one species. Inoculated strains could be detected at a cell concentration of 104 CFU g-1 soil. The analysis of Sphingomonas community structures of several PAH-contaminated soils with the new PCR-DGGE method revealed that soils containing highest phenanthrene concentrations showed lowest Sphingomonas diversity. Sequence analysis of cloned PCR products amplified from soil DNA revealed new 16S rRNA gene Sphingomonas sequences significantly different from sequences from known cultivated isolates, i.e., sequences from environmental clones

- 97 - Chapter 4 grouped phylogentically with other environmental clone sequences available on the web and possibly originated from several potential new species. In conclusion, the new designed Sphingomonas specific PCR-DGGE detection technique successfully analyzed the Sphingomonas communities from polluted soils at the species level and revealed different Sphingomonas members not previously detected with culture- dependent detection techniques.

INTRODUCTION

The Sphingomonas genus was proposed in 1990 (Yabuuchi et al. 1999) to describe a group of bacterial strains isolated from human clinical specimens and hospital environments. During the past decennium Sphingomonas strains have also been isolated from a variety of antropogeneous contaminated environments, including terrestrial (subsurface soil (Mueller et al. 1990; Feng et al. 1997; Lloyd-Jones et al. 1997; Bastiaens 1998; Adkins 1999; Cassidy et al. 1999; Meyer et al. 1999; Momma et al. 1999; Bastiaens et al. 2000; Pinyakong et al. 2000; Sorensen et al. 2001) and rhizosphere soil (Daane et al. 2001)), sediment (river sediments, subsurface sediments (Fredrickson et al. 1995; Fredrickson et al. 1999)) or aquatic habitats (waste water (Coughlin et al. 1999; Meyer et al. 1999; Fujii et al. 2000), ground water (Tiirola et al. 2002), fresh water (Wittich et al. 1992; Stolz 1999; Tabata et al. 1999; Schweitzer et al. 2001), marine water (Gilewicz et al. 1997)), and were shown to posses unique abilities to degrade a variety of pollutants including azo dyes (Stolz 1999), chlorinated phenols (Cassidy et al. 1999; Crawford et al. 1999), dibenzofurans (Keim et al. 1999; Wittich et al. 1999), insecticides (Nagata et al. 1999) and herbicides (Adkins 1999; Kohler 1999). In addition, Sphingomonas strains are often isolated from contaminated soils as degraders of polycyclic aromatic hydrocarbons (PAHs) (Mueller et al. 1990; Khan et al. 1996; Bastiaens et al. 2000; Pinyakong et al. 2000). PAHs are very hydrophobic toxic chemicals with low solubility in water making them poorly available for natural bacterial degradation. Due to their ubiquitous distribution and their diverse catabolic capabilities towards recalcitrant organic pollutants, Sphingomonas strains are thus interesting biocatalysts for soil bioremediation.

- 98 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

Therefore, it is of major interest to be able to monitor the presence, biodiversity and dynamics of Sphingomonas species in the environment. However, until today only a limited number of studies reported on Sphingomonas specific detection and monitoring techniques. Culture-independent molecular identification methods described so far were based on the extraction of typical sphingolipids (Leung et al. 1999) or ribosomal DNA/RNA as marker molecules (Van Elsas et al. 1998; Leung et al. 1999; Thomas et al. 2000; Schweitzer et al. 2001). Several rRNA gene-targeted fluorescent labeled oligonucleotide probes were developed (i) by Thomas et al. (Thomas et al. 2000) to specifically monitor the inoculated PAH-degrading Sphingomonas sp. strain 107 in soil via flow cytometry, and (ii) by Schweitzer et al. (Schweitzer et al. 2001) to analyze the composition of lake aggregate-associated Sphingomonas communities via fluorescent in situ hybridization (FISH). However, sphingolipid analysis gives no information on Sphingomonas diversity and the currently available probes for detection of Sphingomonas by flow cytometry and FISH are detecting all species or some species. Other researchers reported the application of specific PCR to detect Sphingomonas in environmental samples using the 16S rRNA gene as target molecule. A specific primer set and internal probe targeting the ribosomal 16S rRNA genes were designed to monitor specifically by PCR Sphingomonas chlorophenolica RA2 (DSM 8671) seeded in soil (Van Elsas et al. 1998). Two degenerated 16S rRNA gene primer sets (SPf-190/SPr1-852) were designed for PCR-detection of a spectrum of different Sphingomonas species in soil (Leung et al. 1999). However, none of the primer sets so far developed for PCR detection were designed to cover the total Sphingomonas genus and degeneration made them unsuitable to directly assess the diversity of Sphingomonas species in soil using a fingerprinting method like Denaturing Gradient Gel Electrophoresis (DGGE). This paper describes the design of a 16S rRNA gene based non-degenerated primer set selective for specific PCR-detection of all known Sphingomonas species and allowing subsequently differentiation between Sphingomonas species using DGGE analysis. The PCR-DGGE method was used to asses the diversity of the indigenous Sphingomonas community in different PAH-contaminated soils.

- 99 - Chapter 4

MATERIALS AND METHODS

Bacterial strains and culture media. The bacterial strains used in this study are described in Table 4- 1. For genomic DNA-extraction, all strains were cultivated in 869-broth (Mergeay et al. 1985). For evaluation of the method sensitivity, appropriate Sphingomonas strains were cultivated in a phosphate buffered minimal liquid medium previously described (Wick et al. 2001), supplemented with 2 g l-1 of the appropriate PAH compound (ACROS Organics, Geel, Belgium) provided as sole carbon and energy source. All cultures were incubated in the dark on an orbital horizontal shaker at 200 rpm at a constant temperature of 30 °C.

TABLE 4-1 BACTERIAL STRAINS USED IN THIS STUDY

Accession n° of Sphingo108f & Organism (Origin or reference) Capabla of degradingf 16S rRNA gene Sphingo420r$ Class of α–PROTEOBEACTERIA, α-4-Subclass Sphingomonadaceae family Sphingomonas genus Sphingomonas adhaesivae Op-55 (DSM 7418T) NR D16146 + Sphingomonas’ agrestis’ HV3 (Yrjala et al. 1998) nap Y12803 + Sphingomonas aromaticivorans F199 (DSM 12444T) nap, tol, xyl, bip, flu, dibt, cres AB025012 + Sphingomonas asaccharolytica Y-345 (DSM 10564T) NR Y09639 + Sphingomonas capsulata 28 (DSM 30196T) NR D16147 + Sphingomonas chlorophenolica (DSM 7098 T) PCP, TiCP X87161 + Sphingomonas chlorophenolica RA2 (DSM 6824) PCP X87164 + Sphingomonas sp. VM0440 (Springael et al. unpublished) phe AY151392 + Sphingomonas sp. LB126 (Bastiaens et al. 2000) flu AF335501 + Sphingomonas sp. VM0506 (Springael et al. unpublished) flu AF335468 + Sphingomonas sp. LH227 (Bastiaens et al. 2000) phe AY151393 + Sphingomonas macrogoltabida 203 (DSM 8826T) PEG D13723 + Sphingomonas mali Y-347 (DSM 10565T) NR Y09638 + Sphingomonas notatoria UQM2507 (DSM 3183T) NR AB024288 + Sphingomonas parapaucimobilis OH3607 (DSM 7463T) NR D13724 + Sphingomonas paucimobilis KS0301 (LMG2239) NR D38420 + Sphingomonas paucimobilis CL1/70 (DSM 1098T) NR D13725 + Sphingomonas pruni Y-250 (DSM 10566 T) NR Y09637 + Sphingomonas rosa R135 (DSM 7285T) NR D13945 + Sphingomonas sanguis KM2397 (LMG2240T) NR D13726 + Sphingomonas sp. EPA505 (DSM 7526) flu, nap, phe, ant, bflu U37341 + Sphingomonas subartica KF1 (DSM 10700T) TeCP, TiCP X94102 + Sphingomonas subartica KF3 (DSM 10699) TeCP, TiCP X94103 + Sphingomonas sp. LH128 (Bastiaens 1998) phe AY151394 + Sphingomonas suberifaciens CR-CA1 (DSM 7465T) NR D13737 + Sphingomonas terrae (LMG10924) NR D38429 + Sphingomonas terrae E-1-A (DSM 8831T) PEG D13727 + Sphingomonas trueperi (DSM 7225T) NR X97776 + Sphingomonas ursincola KR-99 (DSM 9006T) NR AB024289 + Sphingomonas wittichii RW1 (DSM 6014T) dbf AB021492 + Sphingomonas xenophaga BN6 (DSM 6383 T) 2-nap-sulfonate X94098 + Sphingomonas yanoikuyae AB1105 (DSM 7462T) NR D16145 + Sphingomonas yanoikuyae B1 (DSM 6900) tol, xyl, bip, nap, phe X94099 + Sphingomonas yanoikuyae Pn4S (LMG3925) NR D13946 + Other Sphingomonadaceae genera Porphyrobacter neustonensis (DSM 9434T) NR AB033327 + Porphyrobacter tepidarius OT3 (DSM 10594T) NR AB033328 + Erythrobacter litoralis T4 (DSM 8509T) NR AB013354 + Erythromicrobium ramosum E5 (DSM 8510T) NR AB013355 + Zymomonas mobilis subsp. paniaceae I (LMG448T) NR AF281032 + Other α–Proteobacteria Phyllobacterium rubiacearum (DSM 5893T) NR D12790 - Agrobacterium luteum A61 (DSM 5889T) NR NR - Rhizobium radiobacter L624 (DSM 30147T) NR AJ389904 - Rhizobium radiobacter B6 (DSM 30205) NR D14500 (+) Rhizobium radiobacter B2326 (DSM 30203) NR D14506 (+) Rhizobium rubi TR3 (DSM 6772T) NR D12787 (+) Sinorhizobium meliloti 3DOa2 (DSM 30135T) NR D14509 - Rhodobacter sphaeroides ATH2.4.1 (DSM 158 T) NR D16425 -

- 100 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

Rhodobacter sphaeroides (DSM 160) NR NR - Rhodobacter capsulatus (ATCC 23782) NR NR - Rhodospirillum rubrum B-280 (ATCC 19613) NR NR - Rhodospirillum rubrum S1H (ATCC 25903) NR NR - Brevundimonas diminuta 342 (DSM 7234T) NR AJ227778 - Brevundimonas diminuta PCI818 (DSM 1635) NR X87274 - β - γ- δ -PROTEOBACTERIA Ralstonia metallidurans CH34 (DSM 2839T) NR Y10824 - Burkholderia sp. JS150 (DSM 8530) ben AF262932 - Aeromonas enteropelogenes J11 (DSM 6394T) NR X71121 - Acinetobacter calcoaceticus 46 (DSM 30006T) NR AJ247199 - Pseudomonas putida (DSM 8368) nap, phe, flu, fan NR - Desulfobacter latus AcRS2 (DSM 3381T) NR AJ441315 - Desulfonema magnum 4be13 (DSM 2077T) NR U45989 - Desulfobulbus rhabdoformis M16 (DSM 8777T) NR U12253 - Gram positive BACTERIA Arthrobacter sulfureus 8-3 (DSM 20167T) NR X83409 - Dietzia maris IMV 195 (DSM 43627T) NR X79290 - Mycobacterium frederiksbergense FAn9 (DSM 44346T) fan, phe, pyr AJ276274 - T = species type strain and/or genus type species. NR = Not Reported nap = naphthalene; fan = fluoranthene; pyr = pyrene; flu = fluorene; phe = phenanthrene; ant = antracene; bflu = benzo(b)fluorene; dibt= dibenzothiophene; dibf = dibenzofurane; bip = biphenyl; ben = benzene; tol = toluene; xyl = xylene; TiCP = trichlorophenol; TeCP= tetrachlorophenol; PCP = pentachlorophenol; PEG = polyethylene glycol; cres= cresol. * result of PCR with primers Sphingo108f & GC40-Sphingo429r on pure strain DNA extract: + = high concentration of PCR product, (+) = low concentration of PCR product - = no detectable PCR product

Soil Samples. Soil samples were taken from different historically PAH-contaminated industrial sites and their characteristics are summarized in Table 4-2. The methods applied for chemical and physical analysis have been reported previously (Chapter 2).

TABLE 4-2 CHARACTERISTICS OF SOIL SAMPLES USED IN THIS STUDY

Soil Origin Soil pH TOC PAH conc. Min. oil conc. DNA conc. * PCR† Estimated type (%) (mg kg-1) (mg kg-1) (µg g-1) cell conc.‡ (Cells g-1)° K3840 Gasoline station site (Denmark) sand 8.2 0.50 20 98 2.75 +1/100 106 B101 Coal gasification plant (Belgium) sand 7.0 2.63 107 70 27.25 +1/10 105 TM Coal gasification plant (Belgium) sand 8.0 3.85 506 4600 4.75 +1/100 106 Barl Coal gasification plant (Germany) gravel 8.9 4.63 1029 109 6.15 +1/100 106 AndE Railway station site (Spain) clay 8.1 2.35 3022 2700 3.40 +1/100 106 ND = not determined * DNA recovery per g soil, mean value of 2 parallel extractions of 1 soil sample. † PCR = result of PCR with Sphingo108f & GC40-Sphingo420r on soil DNA extract + = PCR product, 1/10 or 1/1000 = highest template dilution which was still positive in PCR ‡ Cell Conc. = roughly estimated Sphingomonas cell concentration based on a ‘dilution to extinction’ PCR approach

Design of a new Sphingomonas specific 16S rRNA gene primer set. Circa 215 sequences (min 1200bp long) of both environmental and clinical Sphingomonas species available from the GenBank database (Benson et al. 2003) were selected and aligned using the RPDII Hierarchy Browser program (Cole et al. 2003) and the ClustalW software (Higgins et al. 1994). The multiple alignment was further analyzed by the TreeTop software (Brodsky et al. 1995) for phylogenetic tree prediction and with the PLOTCON program (EMBOSS software, Version 2.3.1) to identify variable gene regions. The sequence similarity was calculated by moving a window of 4 bp along the aligned sequences. Within the window, the similarity of any one position was taken to be the average of all the possible pair wise scores (taken from the specified similarity matrix of the imported alignment) of the bases at that position. The average of the position similarities within the window was plotted, resulting in a similarity plot. The primers had to be located in a conserved region and had to amplify a variable

- 101 - Chapter 4 region of maximum 500 bp to allow good DGGE analysis of the amplicons. Several possible primer combinations were visually selected from the constructed alignment of rrn genes of Sphingomonas species. The primer pairs were identified based on selectivity analysis using the Advanced Blast Search program (Genbank, NCBI) (Altschul et al. 1990) and the Sequence Match program (RDP II) (Cole et al. 2003). The final primer set consisted of the forward primer Sphingo108f (5'- GCGTAACGCGTGGGAATCTG -3', E. coli position 108 to 128) and the reverse primer Sphingo420r (5'-TTACAACCCTAAGGCCTTC-3', E. coli position 420 to 401). A 40 basepair long GC-clamp (CGC GGG CGG CGC GCG GCG GGC GGG GCG GGG GCG CGG GGG G) (Muyzer et al. 1993) was attached to 5’ end of the reverse primer to allow DGGE analysis of the amplicons. This new primer couple Sphingo108f and GC40-Sphingo420r amplified a 312 bp sequence of the 16S rRNA gene resulting in a PCR-product of 352 bp long. DNA-extraction. DNA was extracted from cultures and soil as described previously (Chapter 2). The DNA-concentrations in the 100 µl cell extracts and 50 µl soil extracts were measured spectroscopically. For PCR purposes the concentration of pure strain DNA was adjusted to a final concentration of 100 ng µl-1. For Sphingomonas cells, 100 ng of DNA corresponds to circa 2.9×107 cell equivalents and 2.9×107 copies of PCR targets assuming a genomic molecular weight of 3.2 mega base (i.e. ca. 2.1×109 Daltons = 3.5 fg DNA) per cell (Eguchi et al. 2001) and only one 16S rRNA gene copy per genome (Fegatella et al. 1998; Tiirola et al. 2002). To assure that the soil DNA was of good quality for PCR, dilution series of all soil DNA extracts were tested in PCR with universal eubacterial 16S rRNA gene primer pair GC-63f and 518r with the forward primer linked to a 40 bp long GC-clamp (Muyzer et al. 1993). Dilutions of 1:10, 1:100 and 1:1000 soil DNA-extracts in water were further used as template in a dilution-to-extinction PCR with the appropriate primer sets. PCR reaction. PCR reactions with universal eubacterial 16S rRNA gene primers were performed according to the instruction of the authors (Muyzer et al. 1993; Marchesi et al. 1998). The PCR protocol used with the Sphingo108f and GC40-Sphingo420r primer pair consisted of a short denaturation of 15 s at 95°C, followed by 50 cycles of denaturation for 3 s at 95°C, annealing for 10 s at 62°C and elongation for 30 s at 74°C. The last step included an extension for 2 min at 74°C. PCR was performed on Biometra (Biometra, Göttingen, Germany) or Perking Elmer (Perking Elmer, Connecticut, USA) PCR-machines. PCR mixtures contained 100 ng of pure strain DNA or dilutions of soil DNA as templates, 1 U Taq polymerase, 25 pmol of the forward primer, 25 pmol of the reverse primer, 10 nmol of each dNTP, and 1 × PCR buffer in a final volume of 50 µl. The Taq polymerase, dNTPs and PCR buffer were purchased from TaKaRa. DGGE analysis. The PCR-products were checked on 1.5 % agarose gels (MetaPhor, BioWhittaker, Labtrade Inc., Miami, Florida, USA) and directly used for DGGE analysis on polyacrylamide gels as previously described (Muyzer et al. 1998b). Optimal denaturing conditions were defined based on the theoretical melting temperatures of amplification fragments produced with the Sphingo-primer set as calculated with the DAN program (EMBOSS, Version 2.3.1) and the Melt program (Version 1.0.1, INGENY International BV, Goes, The Netherlands). A 6 % percent polyacrylamide gel with a denaturing gradient of 40 % to 75 % (where 100% denaturant gels containing 7 M urea and 40 % formamide) was used for DGGE-analysis. Electrophoresis was performed at a constant voltage of 130

- 102 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

V for 16 h 40 m in 1 × TAE running buffer at 60 °C in the DGGE-machine (INGENYphorU-2, INGENY International BV). After electrophoresis, the gels were stained with 1 × SYBR Gold nucleic acid gel stain (Molecular Probes Europe BV, Leiden, The Netherlands) and photographed under U.V. light using the digital camera system Image Master VDS (Liscap Image Capture software, Version 1.0, Pharmacia Biotech, Cambridge, England). Photofiles were processed and analyzed with the Bionumerics software (Version 2.50, Applied Maths, Kortrijk, Belgium). Sensitivity of PCR detection. To examine the sensitivity of the PCR method to detect Sphingomonas strains in soil, a standard of living cells of Sphingomonas sp. strain LB126 were added at different final cell concentrations (i.e. approximately 105, 104, 103, 101, 100 CFU g-1) to an uncontaminated model soil prior to DNA-extraction. Before they were added to the soil samples, the cultures were filtered over glass wool to remove the excess of PAH crystals and cells were washed twice and finally appropriately diluted in an isotonic aqueous solution of 0.85 % w v-1 NaCl. The total soil DNA extract was subsequently used as template in PCR with the Sphingo-primers and PCR-products were analyzed by DGGE. PCR-DGGE analysis of Sphingomonas communities in PAH-contaminated soils. To assess the presence of Sphingomonas strains in a set of contaminated soils, soil DNA-extracts were analyzed in PCR with the Sphingo-primer set. To roughly estimate the concentration of the detected Sphingomonas cells, dilution series of non-inoculated soil DNA extracts (1:1, 1:10, 1:100 and 1:1000 dilutions in water) were tested in a ‘dilution to extinction’ PCR-approach, similar to the MPN-PCR approach. The final cell density within a soil was deduced from the highest template dilution for which still a PCR product was detected, taking into account that the highest dilution giving a signal contained a cell density approaching the determined detection limit. Parallel soil samples with added cells were regarded as positive PCR controls to assure that negative PCR results with samples without added cells were not due to PCR inhibition effects. 16S rRNA gene amplicons resulting from PCR with the Sphingo-primer set on the soil DNA extracts were cloned into plasmid vector pCR2.1-TOPO using the TOPO Cloning Kit (N.V. Invitrogen SA, Merelbeke, Belgium) as described within the kits protocol without prior concentration or purification. Clones containing recombinant vectors with the appropriate 16S rRNA gene fragment were compared with the soil Sphingomonas community fingerprints using DGGE to identify which bands from the pattern were selected. A selection of clones with different DGGE-patterns was sequenced by the Westburg Company. The 16S rRNA gene-sequences obtained from the cloned PCR-products were submitted to the 'Chimera Check' program (RDPII) (Cole et al. 2003) to detect possible chimeras that could have been formed during PCR (Maidak et al. 1994). A similarity analysis of the 16S rRNA gene sequences was obtained by using the Advanced Blast Search program (Genbank, NCBI) (Altschul et al. 1990). To study the evolutionary relationships between the 16S rRNA gene sequences retrieved from PCR amplified soil DNA and from known Sphingomonas species, clone sequences were imported into the alignment and edited manually to remove nucleotide positions of ambiguous alignment and gaps. Sequence similarities were calculated for the total length of the 16S rRNA gene sequences and corrected using Kimura's two-parameter algorithm to compensate for multiple nucleotide exchange and a distance-based evolutionary tree was constructed using Kimura's corrected similarity values in the Neighbor-Joining algorithm (Saitou et al. 1987). The

- 103 - Chapter 4 topography of the branching order within the dendrogram was evaluated by using the Maximum- Likelihood and the Maximum-Parsimony character-based algorithms in parallel combined with bootstrap analysis with a round of 500 reassemblings. The 16SrRNA gene sequence from some closely related genera from the Sphingomonadaceae family (Zymomonas, Porphyrobacter, Erythrobacter, Sandaracinobacter, etc.) and some more distant related α-Proteobacteria (Rhizobium, Rhodospirillum, Rhodobacter, Sinorhizobium, etc.) were included as an out-group to root the tree. Clone nucleotide sequence accession numbers. The 16S rRNA gene clone sequences retrieved from contaminated soils with the Sphingo-primer set are available from GenBank under accession numbers AY335445 to AY335484.

RESULTS

Design of a Sphingomonas genus specific primer set. The rrn gene is moderately conserved within the Sphingomonas genus as was indicated by a similarity plot created from an alignment of Sphingomonas 16S RNA gene sequences (min. 1300bp long). The alignment showed a minimum similarity of ca. 89% over the total length of the rrn gene within the Sphingomonas genus (data not shown). From the alignment we selected a new non-degenerated primer set that would anneal to 16S rRNA gene sequences and that spanned a region between 200bp and 600bp long with high variability in order to allow differentiation of the various species by DGGE-analysis of the PCR-products. Blast (NCBI) and Sequence Match (RPDII) analysis (April 2003) was used to check primer selectivity. Out of the 6 different primers selected and tested in different appropriate combinations (data not shown), the primer couple Sphingo108f and Sphingo420r was the best combination possible, targeting as many Sphingomonas species as possible and as less as possible non-Sphingomonas sequences. The forward primer Sphingo108f was highly selective for the Sphingomonas genus (Figure 4-1). Of all sequences available in the NCBI database (Cole et al. 2003), currently holding circa 375 Sphingomonas genus sequences of all length, ca. 350 sequences were found 100% homologous to the Sphingo108f primer sequence using the Sequence Match software (RDPII).

- 104 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

Organism(Accession N°) * Primers Sphingo108f Sphingo420r (E.coli 108 -128) (E. coli 420-401) 5' - GCGTAACGCGTGGGAATCTG - 3' 5' - TTACAACCCTAAGGCCTTC - 3' Sphingomonas genus strains S. wittichii DSM 6014T (AB021492) ------S. pituitosa DSM 13101T (AJ243751) ------S. trueperi DSM 7225T (X97776) ------S. paucimobilis DSM 10987T (U37337) ------G------S. parapaucimobilis DSM 7463T (D13724) ------G------S. sanguinis LMG 17325T (D13726) ------G------S. aquatilis IFO 16772T (AF131295) ------S. echinoides DSM 1805T (AB021370) ------S. adhaesiva DSM 7418T (D16146) ------G------S. pruni DSM 10566T (Y09637) ------S. mali DSM 10565T (Y096368) ------S. asaccharolytica DSM 10564T (Y09639) ------S. suberifaciens DSM 7465T (D13737) ------S. yanoikuyae DSM 7462T (D16145) ------S. xenophaga DSM 6383T (X94098) ------S. chlorophenolicum DSM 7098T (X87161) ------S. chungbukensis JCM 11454T (AF159257) ------S. herbicidivorans DSM 11019T (AB042233) ------S. cloacae JCM 10874T (AB040739) ------S. rosa DSM 7285T (D13945) ------S. stygia CIP 10514T (AB025013) ------S. subterranea CIP 105153T (AB025014) ------S. aromaticivorans DSM 12444T (AB025012) ------S. capsulatum DSM 30196T (D16147) ------S. terrae DSM 8831T (D13727) ------G------S. macrogoltabida DSM 8826T (D13723) ------G------S. alaskensis DSM 13593T (Z73631) ------S. taejonensis JCM 11457T (AF131297) ------G------S. subartica DSM 10700T (X941025) ------Other Sphingomonadeaceae family strains Sandaracinobacter sibericus RB16-17 (Y10678) ------G------Zymomonas mobilis LM G448T (AF281032) ------Porphyrobacter tepidarius DSM 10594T (AB033328) ------Porphyrobacter neustonensis DSM 9434T (AB033327) ------Erythrobacter longus DSM 6997T (M59062) ------Erythromicrobium ramosum DSM 8510T (AB013355) ------Non-Sphingomonadaceae strains Rhizobium rubi IFO13261 (D14503) A------A ------Rhizobium rubi DSM 9772T (X67228) A------A ------Rhodobacter sphaeroides 2.4.1T (X53853) A------CG------Methylobacterium radiotolerans JCM 2831T (D32227) A------CG------Rhizobium radiobacter DSM 30147T (AJ389904) A------CA-A ------Methylobacterium organophilum JCM 2833T (D32226) A- - - - - A------CG-A ------Rickettsia massiliae Mtu1T (L36214) A- - - - - A------A ------Rickettsia honei RBT (U17645) A- - - - - A------A ------Bradyrhizobium japonicum DSM 30131T (U69638) A------CG-A ------G------Rhodospirillum rubrum ATCC 11170T (D30778) A- - - - - A------G-A ------G------Caulobacter vibroides CB2AT (M83799) A- - - - - A------CG------T- - - - A------Pseudomonas aeruginosa LMG1242T (Z76651) A- - - - T- C- A------C- - - - - T- - - - A------Pseudomonas putida DSM 291T (Z76667) A- - - - T- C- A------T- - - - A------

* Accession No of 16S rRNA gene sequence in the Genbank (NCBI)

FIGURE 4-1 DNA-SEQUENCE HOMOLOGY BETWEEN THE SPHINGOMONAS GENUS SPECIFIC PRIMERS AND THE 16S RRNA GENE SEQUENCE OF SOME REFERENCE SPHINGOMONAS STRAINS Results are presented in a consensus table of matches. Dashes indicate homologous sequences.

- 105 - Chapter 4

Besides within Sphingomonas strains, the forward primer was also 100% conserved in 16S rRNA gene sequences of Sandaracinobacter, Zymomonas, Porphyrobacter, Erythrobacter or Erythromicrobium strains, which as Sphingomonas belong to the Sphingomonadaceae family (Figure 4-1). Only a few of the sequences with 100% homology to primer Sphingo108f (ca. 20) corresponded to some Caulobacter, Pseudomonas or Rhizobium strains. At least 2 mismatches were found between the primers in 16S rRNA gene sequences of other strains not belonging to the Sphingomonadaceae family (Figure 4-1). The reverse primer Sphingo420r proved to be more conserved, i.e. at least 1600 sequences in the bacterial ribosomal database showed 100% similarity with the primer sequence. Sequences of all genera of the Sphingomonadaceae family, i.e., Sphingomonas, Zymomonas, Porhphyrobacter, Erythrobacter and Erythromicrobium, aligned perfect with the reverse primer sequence. Some Sphingomonas and Sandaracinobacter species had a single mismatch with the reverse primer. Most non-Sphingomonadaceae sequences with 100% homology to the Sphingo420r primer belonged to some strains of the genera Rhizobium, Methylobacterium and Rickettsia. The newly developed Sphingo108f & GC40-Sphingo420r primer couple produced only products of the appropriate size and only with the DNA obtained from all the 34 tested Sphingomonas strains representing different species (Table 4-1), while the other tested primer combinations did not. As expected, positive PCR results also were obtained for most of the test strains belonging to the other Sphingomonadaceae genera, i.e., Porphyrobacter, Erytrobacter, Zymomonas, and Erythromicrobium and faint signals for some Rhizobium strains. In PCR with the DNA of the 11 tested non-α-Proteobacteria genera (Table 4-1) no products were detected. It can thus be concluded that the new designed primer set Sphingo108f and Sphingo420r is selective for the detection of Sphingomonas strains and all bacteria belonging to the Sphingomonadaceaea family.

DGGE-analysis of pure strain PCR fragments amplified with the Sphingo- primer set. In order to examine if DGGE-analysis would allow direct differentiation of Sphingomonas species in mixed environmental communities, a GC40-clamp was attached to the reverse primer Sphingo420r and the PCR obtained 16S rRNA gene fragments were loaded on a DGGE-gel (Figure 4-2).

- 106 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

All tested Sphingomonas stains were characterized by a DGGE-profile consisting of 1 single band, except for S. trueperi DSM 7225T (lane 27) and S. paucimobilis DSM 7463T (lane 20) that showed 2 less intense additional bands. Strains which are very closely related based on 16S rRNA gene, most likely belonging to the same species, showed identical DGGE-fingerprints as indicated for the two S. chungbukensis strains (lanes 1 and 2) or three S. subartica strains (lanes 21, 22 and 23). Different species showed mostly different DGGE fingerprints. However, some very closely related species (amplicon similarity > 97%) displayed similar DGGE-fingerprints like for example S. paucimobilis and S. parapaucimobilis (lanes 24 and 25) or S. asaccharolytica and S. pruni (lanes 9 and 10). Similar DGGE-fingerprints were also found for 2 more distantly related species such as S. mali and S. terrae (lanes 5 and 6).

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

FIGURE 4-2 SPHINGOMONAS SPECIES DIFFERENTIATION BY DGGE-ANALYSIS OF DNA- FRAGMENTS AMPLIFIED WITH PRIMERS SPHINGO108F & GC40-SPHINGO420R. The separate lanes represent the different species specific DGGE melting profiles of different tested Sphingomonas strains. Lanes: 1, Sphingomonas sp. VM0506; 2, Sphingomonas sp. LB126; 3, S. macrogotabida DSM 8826T; 4, S. notatoria DSM 3183T; 5, S. mali DSM 10565T; 6, S. terrae DSM 8831T; 7, S. yanoikuyae DSM 7462T; 8, S. suberifaciens DSM 7465T; 9, S. asaccharolytica DSM 10564T; 10, S. pruni DSM 10566T; 11, S. capsulata DSM 30196T; 12, S. rosa DSM 7285T; 13, S. aromaticivorans DSM 12444T; 14, S. xenophaga DSM 6383T; 15, Zymomonas mobilis LMG448T; 16, Erythrobacter litoralis DSM 8509T; 17, S. sp. LH227; 18, S. wittichii DSM 6014T; 19, Sphingommonas sp. EPA505; 20, S. paucimobilis DSM 1098T; 21, Sphingomonas sp. LH128: 22, S. subartica DSM 10700T; 23, S. subartica DSM 10699; 24, S. paucimobilis LMG2239; 25, S. parapaucimobilis DSM 7463T; 26, S. sanguis LMG2240, 27, S. trueperi DSM 7225T; 28, S. flava DSM 6824; 29, S. adhaesiva DSM 7418T. Lanes were ordered using the Bionumerics software to group and compare several DGGE-profiles.

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Limit of detection of Sphingomonas in soil using the PCR protocol with primers Sphingo108f & GC40-Sphingo420r. An inoculated soil experiment was set up to investigate the amplification sensitivity of the new primer set Sphingo108f and GC40-Sphingo420r. Living cells of Sphingomonas sp. LB126 were added at different final cell concentrations to an uncontaminated model soil prior to DNA-extraction. Sphingomonas strain LB126 could be detected down to a cell concentration of 2x104 CFU g-1.

Analysis of Sphingomonas soil communities with primer set Sphingo108f & GC40-Sphingo420r. Different PAH-contaminated soil samples with different contamination records from different European sites (Table 4-2) were screened for the presence of Sphingomonas species by means of PCR with the Sphingo-primer set on total soil DNA-extracts followed by DGGE-analysis of the resulting 16S rRNA gene amplicons for diversity analysis. The DNA-concentration in the soil extract indicated an approximate DNA recovery of 0.135 to 1.375 µg DNA g-1 soil. Assuming that 100 % of the in situ biomass represents bacteria and a bacterial cell contains in general 5 fg of DNA per cell (Chandler et al. 1999) this would theoretically be equivalent to 2.7x107 to 2.8x108 cells g-1 soil. Endogenous Sphingomonas could be detected in all tested soils (Figure 4-3). The ‘dilution to extinction’ PCR method estimated roughly the total Sphingomonas cell concentration between 105 to 106 cells per gram of soil (Figure 4- 2). The DGGE profiles of the Sphingomonas community in the soil samples retrieved by PCR with primer set Sphingo108f & GC40-Sphingo420r were relatively complex, comprising several bands for each sample (Figure 4-3). Soils containing highest concentrations of PAHs showed the lowest number of Sphingomonas 16S rRNA gene bands while less contaminated soils showed a significantly higher number of bands in DGGE fingerprinting. The diversity differences among the samples were further analyzed by random cloning of 16S rRNA gene PCR-products and sequencing of clones showing diverse DGGE-patterns. A comparison of the soil DGGE profiles and the DGGE profiles obtained with the soil clones allowed presumptive identification of some bands (Figure 4-3).

- 108 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

Higher PAH conc. Lower PAH conc.

AndE Barl TM B101 3840

TM/11

LB126 B101/1 TM/1 B101/2 T Barl/9 DSM7465 TM/3 Barl/3 TM/9 Barl/5 AndE/1 Barl/4 B101/3 3840/2 AndE/2 Barl/7 TM/10 AndE/6 Barl/2 3840/3 LH128 TM/13 B101/6 AndE/5 Barl/6 3840/5 3840/4 AndE/4 Barl/8 TM/12 B101/4 AndE/3 TM/8 Barl/1 3840/1 TM/6 B101/7 TM/5 TM/2 TM/7 B101/5 TM/4

FIGURE 4-3 DGGE ANALYSES OF INDIGENOUS SPHINGOMONAS COMMUNITIES IN NATURAL SOIL SAMPLES USING PRIMERS SPHINGO108F & GC40-SPHINGO420R IN PCR. The separated lanes indicate the DGGE-fingerprints of the indigenous Sphingomonas community of PAH-contaminated soils K3840, B101, TM, Barl and AndE. Cloned ‘bands’ are indicated within the soil fingerprint based on the comparison of migration profiles of pure clones and the soil profile. A mix of six strains was used as marker during DGGE-analysis.

Most cloned sequences matched significantly (93-99% similarity) with 16S rRNA gene Sphingomonas sequences from the databases in Blast analysis (Table 4-3). However, 60% of the Blast results were sequences from ‘uncultured’ α-Proteobacteria and Sphingomonas with unknown phylogenetic position within the Sphingomonas genus. To further identify the species lineation, the 40 cloned 16S rRNA gene sequences were aligned with ca. 200 database sequences and a phylogenetic tree was constructed. Phylogenic analysis revealed that all clone sequences exhibited high levels of similarity to sequences typical of the Sphingomonadaceae family, except one (clone Barl/9) that was more related to other α-Proteobacteria (Table 4-3) (Figure 4- 4).

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TABLE 4-3 ANALYSIS RESULTS OF 16S RRNA GENE CLONED SEQUENCES RETRIEVED FROM DIFFERENT SOIL SAMPLES

Soil Clones (Accession N°) Best match in Blast analysis (Altschul et al. 1990) Closest Species match

K3840 3840/1 (AY335480) 91% Uncultured Sphingomonas clone CEA (AF392653) / 3840/2 (AY335481) 95% S. witflariensis W-50 (AJ416410) S. witflariensis 3840/3 (AY335482) 98% Uncultured Sphingomonas clone D104 (AF337854) putative new Sphingomonas species 2 3840/4 (AY335483) 98% Uncultured Sphingomonas clone 367-2 (AF423253) putative new genus 3840/5 (AY335484) 98% Uncultured Sphingomonas clone 739-2 (AF42389) putative new Sphingomonas species-2 B101 B101/1 (AY335454) 97% ‘Afipia genospecies 11‘ (U87782) putative new Sphingomonas species-2 B101/2 (AY335455) 99% Uncultured Sphingomonas clone 768-2 (AF423293) putative new genus B101/3 (AY335456) 96% Sphingomonas sp. K6 (AJ000918) putative new Sphingomonas species-2 B101/4 (AY335459) 95% Sphingomonas sp. SIA181-1A1 (AF395032) / B101/5 (AY335460) 95% Uncultured Sphingomonas clone 739-2 (AF42389) putative new Sphingomonas species-2 B101/6 (AY335457) 97% Uncultured Sphingomonas clone BIccii3 (AJ318120) putative new Sphingomonas species-3 B101/7 (AY335458) 98% Sphingomonas sp. RSI-28 (AJ252595) putative new Sphingomonas species 2 TM TM/1 (AY335468) 96% Uncultured Sphingomonas clone WD290 (AF058299) S. wittichii TM/2 (AY335476) 96% Uncultured Sphingomonas clone TRS1 (AJ006014)) Sandaracinobacter sibericus TM/3 (AY335470) 98% Porphyrobacter sp. MBIC3936 (AF058299) Erythrobacter longus TM/4 (AY335479) 96% Uncultured Sphingomonas clone 739-2 (AF42389) putative new Sphingomonas species-2 TM/5 (AY335477) 97% Uncultured Sphingomonas clone WD249 (AJ292599) putative new Sphingomonas species-2 TM/6 (AY335475) 98% Uncultured Sphingomonas clone saf2-409 (AF078258) putative new Sphingomonas species-2 TM/7 (AY335478) 96% Uncultured Sphingomonas clone 739-2 (AF42389) putative new Sphingomonas species-2 TM/8 (AY335474) 96% Sphingomonas sp. KA1 (AB064271) S. subartica TM/9 (AY335469) 99% ‘Afipia genospecies 13‘ (U87784) putative new genus TM/10 (AY335471) 97% Uncultured Sphingomonas clone t008 (AF422583) S. hassiacum TM/11 (AY335467) 97% Uncultured Sphingomonas clone S23435 (D84626) S. hassiacum TM/12 (AY335473) 98% Uncultured Sphingomonas clone a13104 (AY103311) putative new Sphingomonas species-2 TM/13 (AY335472 ) 97% Uncultured Sphingomonas clone D104 (AF337854) putative new Sphingomonas species 2 Barl Barl/1 (AY335453) 98% Sphingomonas sp. SRS2 (AJ251638) S. wittichii Barl/2 (AY335450) 98% Uncultured Sphingomonas clone AW030 (AF385533) putative new Sphingomonas species-1 Barl/3 (AY335446) 98% S. suberifaciens (D13737) S. suberifaciens Barl/4 (AY335448) 99% Uncultured Sphingomonas clone AW030 (AF385533) putative new Sphingomonas species-1 Barl/5 (AY335447) 97% Uncultured Sphingomonas clone IAFR401 (AF270954) S. suberifaciens Barl/6 (AY335451) 96% Sphingomonas sp. K6 (AJ000918) S. suberifaciens Barl/7 (AY335449) 97% Uncultured Sphingomonas clone IAFR401 (AF270954) S. suberifaciens Barl/8 (AY335452) 97% S. xenophaga UN1F2 (U37346) S. xenophaga Barl/9 (AY335445) 93% Uncultured Sphingomonas clone WD2107 (AJ292610) α-proteobacterium And AndE/1 (AY335461) 95% Uncultured Sphingomonas clone BIccii3 (AJ318120) putative new Sphingomonas species-3 AndE/2 (AY335462) 99% Sphingomonas sp. GTIN11(AY056468) S. cloacae AndE/3 (AY335466) 98% S. xenophaga UN1F2 (U37346) S. cloacae AndE/4 (AY335465) 99% Sphingomonas sp. GTIN11(AY056468) S. cloacae AndE/5 (AY335464 ) 99% Sphingomonas sp. GTIN11(AY056468) S. cloacae AndE/6 (AY335463) 99% Sphingomonas sp. GTIN11(AY056468) S. cloacae

Only a few clone sequences were placed in groups with Sphingomonadaceae genera different from Sphingomonas like Sandaracinobacter (clone TM/2) or Erythrobacter (clone TM/3), that are intermixed with the clusters of the Sphingomonas genus in the phylogenetic tree (Figure 4-4). Thus, most cloned sequences were affiliated with true Sphingomonas sequences, confirming the specificity of the new designed Sphingo- primer set. However, only a very small percentage of cloned sequences (5 out of 40) seemed to be related to cultured PAH-degrading identified Sphingomonas species such as S. wittichii (Barl/1 and TM/1), S. yanoikuyae and S. xenophaga (Barl/8), S. chilensis (3840/2) and S. subartica (Barl/8).

- 110 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils. 14 13 12 11 10 9 8 7 6 5 4 3 2 1 0

Out group 93 Barl/9, 3840/1

72 Sphingomonas sp. Ellin426 (AF432250) TM/9, 3840/4, B101/2 TM/2 100 Sandaracinobacter sibericus RB16-17 (Y10678) S. wittichii DSM6014T (AB021492) * Barl/1, TM/1

93 S. pituitosa DSM13101T (AJ243751) S. treuperi DSM7225T (X97776) S. paucimobilis DSM1098T (U37337) S. parapaucimobilis DSM7463T (D13724) T I 93 85 S. sanguinis LMG17325463 (D13726) S. aquatilis IFO16772T (AF131295) S. echinoides DSM1805T (AB021370) S. adhaesiva DSM7418T (D16146) S. pruni DSM10566T (Y09637) S. mali DSM10565T (Y09638) S. asaccharolytica DSM10564T (Y09639)

93 B101/4 S. ‘aerolata’ NW12 (AJ42940)

80 S. ‘aurantiaca’ MA405 (AJ42938)* S. ‘faenia’ MA-olki (AJ42939) Barl/2, Barl/4 (putative new species 1) Sphingomonas sp. AW30 (AF386533) 77 Sphingomonas sp. QSSC5-6 (AF170744) B101/1, B101/3, B101/5, B101/7, Sphingomonas sp. RSI-28 (AJ252595) 3840/3, 3840/5, TM/4, TM/5, 96 71 Sphingomonas sp. SIA181-1A1 (AF39032) TM/6, TM/7, TM/12, TM/13 Sphingomonas sp. JCM7370 (AF170744) (putative new species 2)

100 Zymomonas mobilis LMG448T (AF281032)

74 Sphingomonas sp. S14 (AY190148) 88 Barl/3, Barl/7, Barl/5, Barl/6 Sphingomonas. sp. K6 (AJ000918) T 59 S. suberifaciens DSM7465 (D13737) 88 96 100 Sphingomonas sp. CF06 (U52146)*

87 IIa S. yanoikuyae DSM7462T (D16145)* 93 T Barl/8 87 S. xenophaga DSM6383 (X94098)*

87 S. chlorophenolicum DSM7098T (X87161)* IIb 93 92 S. chungbukensis JCM11454T (AF159257)* S. herbicidivorans DSM11019T (AB042233)

87

87 Sphingomonas sp. LH227 (AY151393) * S. cloacae JCM10874T (AB040739) AndE/2, AndE/3, AndE/4, AndE/5, AndE/6

100 T 95 S. rosa DSM7285 (D13945) S. stygia CIP10514T (AB025013)* 77 S. subterranea CIP105153T (AB025014)* IIIa 91 S. aromaticivorans DSM12444T (AB025012)* 77 S. capsulata DSM30196T (D16147)

S. terrae DSM8831T (D13727)* T 77 S. macrogoltabida DSM8826 (D13723)* IV T 74 S. alaskensis DSM13593 (Z73631) S. taejonensis JCM11457T (AF131297) S. chilensis DSM14889T (AF367204)* S. ‘witflariensis’ W-50 (AJ416410) 3840/2

79 IIIb T 97 S. subartica DSM10700 (X941025)* TM/8

77 T 83 S. hassiacum DSM14552 (AJ416411) TM/10, TM/11

74 56 Sphingomonas sp. SI-15(AJ252582) And/E, B101/6 (putative new speces 3) 76 Porphyrobacter tepidarius DSM10594T (AB033328) T 84 Porphyrobacter nuestonensis DSM9434 (AB033327) Erythrobacter longus DSM6997T (M59062) TM/3 Erythromicrobium ramosum DSM8510T (AB013355)

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FIGURE 4-4 PHYLOGENETIC ANALYSES OF SPHINGOMONAS SEQUENCES RETRIEVED FROM SOIL DNA EXTRACT USING PRIMERS SPHINGO108F & GC40-SPHINGO420R IN PCR. The phylogenetic relationship of cloned sequences is indicated in a character-based evolutionary tree based on the total length of the 16S rRNA gene sequences and constructed using the Neighbourg Joining algorithm. An out-group of the closely related genera Rhizobium and Rhodospirillum was included to root the tree. The bar at the top indicates the similarity percentage, 1% indicating 1 nucleotide substitution per 100 positions. The tree was tested for branching order confidence by Maximum-Parsimony analysis and a round of 500 bootstraps. Bootstrap values are indicated at branch- points and values above 70% indicating reliable branches. Extended branches were collapsed to form smaller blocks. Most important representative strains are indicated per block with the accession numbers of the sequences indicated between brackets. Species harboring PAH-degrading isolates are indicated with an *. Positions of the clone sequences retrieved from soil are indicated on the right of the tree. Species are grouped based on their 16S rRNA gene sequence similarity. Species groups resembled the clustering previously described (Takeuchi et al. 2001) that divided the Sphingomonas genus in 4 new genera based on the 16S rRNA gene dendrogram. Later, this division of the Sphingomonas genus was reconsidered (Yabuuchi et al. 2002) due to the lack of phenotypic and biochemical evidence. The clusters in the figure indicated as I to IV represent the phylogenetic clusters previously assigned to the genera ‘Sphingomonas sensu stricto’, ‘Sphingobium’, Novosphingobium’ and ‘Sphingopyxis’, respectively (Takeuchi et al. 2001).

These culturable PAH-degrading Sphingomonas isolates are exclusively related to strains found in the former ‘Sphingobium’, ‘Sphingopyxis’ and ‘Novosphingobium’ genera proposed in 2001 (Takeuchi et al. 2001). There were no PAH-degrading isolates or cloned sequences from PAH-contaminated soil found to be related to any of the species of the former ‘Sphingomonas sensu stricto’ genus. Most clone sequences isolated in this study were rather grouped in clusters with other uncultured Sphingomonas 16S rRNA gene sequences and a few unidentified Sphingomonas sp. 16S rRNA gene sequences. Thus, these groups could represent 16S rRNA gene sequences of new (uncultivable) species within the Sphingomonas genus. The cluster with isolate ‘Sphingomonas sp. Ellin4265’ could even represent a new genus within the Sphingomonadaceae family different from the Sphingomonas genus, because of its organization in the phylogenetic tree in a separate branch together with the Sandaracinobacter genus. Other 16S rRNA gene clones were grouped in possibly new Sphingomonas species with (i) isolate Sphingomonas sp. AW030 (species-1), (ii) isolates Sphingomonas sp. SIA181-1A1 and RSI-28 (species-2), or (iii) isolate Sphingomonas sp. SI-15 (species-3). A special high fraction of cloned sequences (12 of 40 clones) was found in the clusters of possible new species-2. Most sequences originating from 1 soil were relatively taxonomically spread over the total Sphingomonas genus. Except for the sequences originating from soil AndE, the most heavily contaminated soil tested, for which 5 out of 6 sequences grouped together in the cluster with S. cloacae IAM14885T.

- 112 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

DISCUSSION

To analyze and monitor the diversity and dynamics of the Sphingomonas population during bioremediation processes, a detection method allowing simultaneous detection of several Sphingomonas was developed. The up to now available primer combinations based on 16S rRNA gene were relative strain and/or species specific (Van Elsas et al. 1998; Leung et al. 1999) and were not suited for simultaneous detection of all PAH-degrading Sphingomonas species. Therefore, we developed a new set of Sphingomonas genus specific 16S rRNA gene primers; primer set Sphingo108f & Sphingo429r. As the primer set had to target the whole Sphingomonas genus, we were not able to exclude the detection of also other Sphingomonadaceae genera such as Zymomonas, Porphyrobacter, Erythrobacter and Erythromicrobium, intermixed with the Sphingomonas genus branches in the 16S rRNA gene based phylogenetic tree of the Sphingomonadaceae family, and some Rhizobium stains. Most tested Sphingomonas species were characterized by a single-band DGGE fingerprint of the amplicon obtained after PCR with the Sphingo108f and GC40- Sphingo420r primer set. A multiple band DGGE-pattern was found for only 2 out of 40 tested strain. A multiple band DGGE-fingerprint for a pure strain could indicate multiple 16S rRNA gene copies with sequences divergence. So far only 2 references could be found that report on the rRNA gene copy number in Sphingomonas species. Both reports show only 1 rrn gene copy number for Sphingomonas strains Sphingomonas sp. MT1 (DSM 13663) (Tiirola et al. 2002) and S. alskensis RB2256 (DSM 13593T) (Fegatella et al. 1998). In addition, also in the draft genome sequence of S. aromaticivorans DSM 12444, available at the JGI website, so far only one 16S rRNA gene copy has been identified in one contig (http://www.jgi.doe.gov/). However, 1 rrn gene copy is relative exceptional in the bacterial world: in most prokaryotes the ribosomal DNA consists of tandem repeated arrays of the rrn genes (Klappenbach et al. 2001). The closely related Zymomonas mobilis ZM4 (ATCC 31821) for example contains 4 gene copies (Kang et al. 1998). Further molecular analysis is needed to confirm that the tested S. trueperi and S. paucimobilis species type strains indeed contains multiple rrn gene copies that could explain the multiple- band DGGE-pattern.

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Pure strain DGGE-fingerprints were mostly ‘inter’ and ‘intra’ species specific, i.e., strains officially belonging to the same species showed identical DGGE-fingerprints and different species showed different DGGE fingerprints. Overlapping fingerprints were found for some strains and species. Similarly, TGGE- and DGGE-analysis of 16S rRNA gene fragments could not discriminate between several species of Burkholderia (Falcão Salles et al. 2002) and Bifidobacterium (Satokari et al. 2001) or Arthrobacter and Nocardioides (Felske et al. 1999), due to the high levels of conservation of the amplified 16S rRNA gene fragments. It is clear that the practical resolution limit of the DGGE-technique is at the species or genus level or intermediate between the two, depending on the gene conservation level within the taxonomic group that is under investigation. However, all currently known species grouping related PAH-degrading Sphingomonas strains could be well separated on DGGE gel indicating that the new developed PCR-DGGE technique was suitable to assess the diversity and dynamics of currently known PAH-degrading Sphingomonas populations in soil. These results suggest that each band in a Sphingomonas community DGGE-fingerprint of environmental samples produced by the Sphingo- primer set would mostly indicate only 1 species or very closely related species. It was proven that the new Sphingomonas specific primer set was still amplifying 16S rRNA gene from different species at cell concentrations of 104 CFU g-1 in different soil types. This detection limit could be expected for all Sphingomonas species since most Sphingomonas species seem to contain only one 16S rRNA gene copy. The same cell concentrations for different species would lead to the same template target concentrations (16S rRNA gene concentration) and thus same detection levels. The detection limit of 104 CFU g-1 is lower than other reported detection sensitivities for similar direct PCR methods, as for example for Burkholderia species (5x105 CFU g-1) (Falcão Salles et al. 2002) or Mycobacterium species (ca. 106 CFU g-1) (Chapter 2). The detection limit of 104 CFU g-1 is especially low, knowing that Sphingomonas species contain probably only one copie of the rrn genes in their DNA. Most other soil bacteria can contain many copies per cell (e.g. Burkholderia 5-6 copies) what will improve the cell detection limit. The good results concerning detection limit obtained using the Sphingo-primer DGGE method are probably contributed to the efficient DNA-extraction and optimized PCR conditions.

- 114 - Occurrence and diversity of Sphingomonas species in PAH-contaminated soils.

Finally, the new developed PCR-DGGE method using the new Sphingo-primer set allowed us to analyze the indigenous Sphingomonas population in 5 different PAH- contaminated soils. Sphingomonas species were found present in all tested soils, originating from very different locations and characterized by very different geological and chemical properties. Their relatively high cell concentrations of 105 to 106 cells per g of soil, and their frequent isolation from contaminated soils during enrichments on PAHs as carbon sources (Mueller et al. 1990; Khan et al. 1996; Bastiaens et al. 2000; Pinyakong et al. 2000) indicate that Sphingomonas strains seem to be important colonizers and possibly endemic pollutant degraders in PAH- contaminated soils. Sequence analysis of DGGE band patterns revealed the presences of ‘new’ 16S rRNA gene sequences which were only limited related to identified species and cultivated PAH-degrading isolates. Although the analysis was only based on relatively short sequences (312 bp), the soil-extracted Sphingomonas sequences indicate the presence of possibly 4 new Sphingomonas species and possibly 1 new Sphingomonadaceae genus. These results were compared with the results obtained for the same soil samples by a previously developed culture-dependent Sphingomonas detection method, i.e., a selective plating technique based on the intrinsic streptomycin resistance and the typical yellow morphotype of Sphingomonas (Vanbroekhoven et al. unpublished). The dominant cultivable Sphingomonas isolated in that work were very different from the dominant Sphingomonas detected by our molecular method. Based on 16S rRNA gene sequence, the isolates were mostly grouped in an unidentified cluster - possibly a new species - with Sphingomonas sp. LH227 strain (Bastiaens et al. 2000) (9 out of 22 isolates) or in a cluster with S. taejonensis, S. chilensis and S. witflariensis (5 out of 22 isolates). Only a very few of our clone sequences were related to 16S rRNA genes of the isolates, and if there was a relationship, clones and isolates seldom originated form the same PAH-contaminated soil. It might be that the dominant strains detected by the PCR-based method are streptomycine sensitive and therefore excluded from the population detected by the culture-dependent approach. However, this is unlikely since all Sphingomonas species tested so far, are streptomycine resistant without exception. Moreover, most of our cloned sequences were most similar to sequences of other uncultured Sphingomonas. Thus, based on the nature of the new sequences detected using the culture-independent technique, these

- 115 - Chapter 4 sequences most likely represent truly ‘non culturable’ Sphingomonas strains present in soil. A diverse group of Sphingomonas strains belonging to different species clusters in the genus was present in low and moderated contaminated soils at relatively equal cell concentrations. Soils containing high concentrations of PAHs (mainly phenanthrene) were characterized with less complex DGGE band patterns than less contaminated soils, and hence seem to be dominated by a less diverse group of Sphingomonas species. Our results may suggest that high PAHs concentrations have enriched a few Sphingomonas strains in a very high concentration which possibly masked the detection of other species that are present in lower concentrations. The soil DGGE- fingerprinting technique did clearly show some additional community information (non-cloned fainter bands in the fingerprints) that simple cloning procedures could not reveal. Pure cloning strategies did not allow a complete qualitative nor an accurate quantitative determination of the microbial population presented by the gene pool extracted from the habitat under study as previously concluded by Liesack et al. (Liesack et al. 1991). More intense bands within the DGGE-fingerprint were clearly cloned more easily. In conclusion, the PCR-DGGE detection method described in this study, based on newly developed Sphingomonas specific primers, proved to be a powerful tool for analyzing the Sphingomonas population diversity and dynamics in environmental samples. Furthermore, the primers developed in this study could be useful in a RT- PCR approach targeting the rRNA in order to identify the active Sphingomonas strains involved in PAH biodegradation in the environment.

ACKNOWLEDGEMENTS

This work was supported by the European Commission, through the funding of the Biovab (EC Contract BIO4-CT97-2015) and Biostimul projects (EC Contract QLRT- 1999-00326). We thank S. Schioetz-Hansen, J. Amor and J. Vandenberghe for providing the soil samples investigated in this study.

- 116 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils.

CHAPTER 5

OCCURRENCE OF SPHINGOMONAS SP. EPA505 RELATED STRAINS IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS).

* REDRAFTED AFTER: LEYS NATALIE, RYNGAERT ANNEMIE, BASTIAENS LEEN, TOP EVA, VERSTRAETE

WILLY, SPRINGAEL DIRK (REVISED) CULTURE-INDEPENDENT DETECTION OF SPHINGOMONAS SP. EPA505

RELATED STRAINS IN SOILS CONTAMINATED WITH POLYCYCLIC AROMATIC HYDROCARBONS (PAHS),

MICROB. ECOL.

ABSTRACT

The Sphingomonas genus hosts many interesting pollutant degrading isolates. Sphingomonas sp. EPA505 is the best studied Sphingomonas strain, capable of degrading polycyclic aromatic hydrocarbons (PAHs) as sole source of carbon and energy. Based on 16S rRNA gene sequence analysis, Sphingomonas sp. strain EPA505 forms a separate branch in the Sphingomonas phylogenetic tree grouping exclusively PAH-degrading isolates. For specific PCR detection and monitoring of PAH-degrading Sphingomonas sp. EPA505 and related strains in PAH-contaminated soils, a new 16S rRNA gene based primer set was developed. The new primer set was shown to be highly selective for Sphingomonas sp. strain EPA505 as it only amplified DNA from strain EPA505 and not from other tested Sphingomonas strains or soil bacteria not belonging to the Sphingomonas genus. Using DNA extracts of a variety of inoculated PAH-contaminated soils, the primer pair was able to detect EPA505 in concentrations as low as 102 cells per gram of soil. In 4 out of 5 tested PAH- contaminated soils, we detected 16S rRNA gene fragments that were 99 to 100% similar to the corresponding gene of strain EPA505. The ubiquitous presence of EPA505 related Sphingomonas strains in PAH-contaminated soils with very different contamination profiles and different origin, suggests an important role of this type of Sphingomonas in the natural Sphingomonas community colonizing PAH- contaminated sites.

INTRODUCTION

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The Sphingomonas genus is known to include many strains that are capable to degrade xenobiotic and recalcitrant pollutants, including polycyclic aromatic hydrocarbons (PAHs) (Mueller et al. 1990; Kästner et al. 1994; Khan et al. 1996; Balkwill et al. 1997; Bastiaens 1998; Bastiaens et al. 2000; Pinyakong et al. 2000), azo dyes (Stolz 1999), chlorinated phenols (Cassidy et al. 1999; Crawford et al. 1999; Fujii et al. 2001), dibenzo- and carbo-furans (Feng et al. 1997; Keim et al. 1999; Wittich et al. 1999), insecticides (Nagata et al. 1999) and herbicides (Weissenfels et al. 1991; Adkins 1999; Kohler 1999; Sorensen et al. 2001). Their frequent isolation from soils and waters contaminated with polycyclic aromatic hydrocarbons (PAHs) indicates that Sphingomonas strains are probably key members of the natural PAH- degrading microflora (Mueller et al. 1990; Weissenfels et al. 1991; Kästner et al. 1994; Balkwill et al. 1997; Bastiaens 1998; Bastiaens et al. 2000; Vanbroekhoven et al. unpublished). Using a culture independent PCR-DGGE fingerprinting technique targeting exclusively the Sphingomonas population, Sphingomonas were indeed found to be ubiquitous in a variety of PAH-contaminated soils from very different origin (Chapter 4). This culture-independent 16S rRNA gene based Sphingomonas detection technique revealed the existence of many new (possible uncultivable) Sphingomonas strains differing from most currently cultured and identified Sphingomonas species (Chapter 4). Based on 16S rRNA gene sequence, most of the cultured PAH-degrading Sphingomonas isolates (Mueller et al. 1990; Weissenfels et al. 1991; Kästner et al. 1994; Balkwill et al. 1997; Bastiaens 1998; Bastiaens et al. 2000) are close relatives of the S. yanoikuyae, S. herbicidivorans and S. chlorophenolica species type strains. These three species cluster phylogenetically in a group formally described as ‘the Sphingobium genus’ (Takeuchi et al. 2001). Over the last years, many new PAH- and pesticide-degrading strains and even new species such as S. xenophaga (Stolz et al. 2000), S. chungbukensis (Kim et al. 2000) or S. cloaca (Fujii et al. 2001) have been added to this ‘Sphingobium’ cluster and have emphasized the environmental importance of this group of Sphingomonas strains. An important PAH-degrading strain from this group is Sphingomonas sp. strain EPA505. Strain EPA505 has been isolated in 1989 from soil contaminated with coal tar creosote in Pensacola, USA (Mueller et al. 1990) and is one of the best described PAH-degrading Sphingomonas strains. EPA505 is known for its wide PAH-degradation versatility and can utilize naphthalene, 1- and 2-methylnaphthalene, 2,3-dimethylnaphthalene, fluoranthene, phenanthrene, anthracene and benzo(b)fluorene as sole source of carbon (Mueller et

- 118 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils. al. 1990; Mueller et al. 1997). The catabolic pathways and aromatic catabolic genes of PAH-degradation of EPA505 have been identified (Story et al. 2000; Story et al. 2001; Pinyakong et al. 2003a) and EPA505 has been often used as model strain to study the effects of surfactants on PAH-degradation (Willumsen et al. 1998a; Willumsen et al. 1998b; Barkay et al. 1999; Osterreicher-Ravid et al. 2000). For biorestauration of PAH-contaminated sites, it is of great interest to study the occurrence and diversity of specific groups of culturable PAH-degrading Sphingomonas such as strain EPA505 in the environment in more detail. Specific detection methods would allow monitoring the survival and activity of such PAH- degrading Sphingomonas species during soil bioremediation processes. Culture- independent PCR-based detection methods would be preferable above culture- dependent methods, as they allow a rapid assessment of both culturable and unculturable Sphingomonas populations in the soil. In addition, unlike probe hybridization based methods such as FISH, PCR-based methods deliver new gene sequence information. In principle, specific species might be recognized in fingerprint patterns obtained by DGGE of 16S rRNA gene amplicons recovered from soil DNA by PCR targeting the whole Sphingomonas genus (Chapter 4). Such method will however only detect the most abundant species. A strain- or species-specific PCR- based detection method is therefore needed to reveal additional information on the Sphingomonas species that are otherwise missed. However, to our knowledge, a molecular method (using a 16S rRNA gene based primer set) to detect a specific Sphingomonas species has only been developed so far for one species, i.e., S. chlorophenolica RA2 (DSM 8671) (Van Elsas et al. 1998). For PAH-degrading Sphingomonas strains there are currently no species or strain specific primer sets available. The purpose of this work was to develop a new specific primer set for selective PCR- based monitoring of PAH-degrading Sphingomonas sp. strain EPA505 and relatives. First, a new 16S rRNA gene based primer set was selected and tested for its specificity and sensitivity. Afterwards the new primers were used to examine the presence of strains related to PAH-degrading Sphingomonas strain EPA505 in a set of PAH-contaminated soils.

MATERIALS AND METHODS

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Bacterial strains and growth conditions. The bacterial strains used in this study are described in Table 5-1. Strains were cultivated in 869-broth (Mergeay et al. 1985). All cultures were grown in the dark on an orbital horizontal shaker at 200 rpm at a constant temperature of 30 °C.

TABLE 5-1 BACTERIAL STRAINS USED IN THIS STUDY

16S rRNA gene Reference or origin Signal in PCR with Organism Accession n° EPAf & EPAr † Class of α–PROTEOBEACTERIA, α-4-

Subclass Sphingomonadaceae family Sphingomonas genus Sphingomonas adhaesivae Op-55 D16146 DSM 7418T - Sphingomonas aromaticivorans F199 AB025012 DSM 12444T - Sphingomonas asaccharolytica Y-345 Y09639 DSM 10564T - Sphingomonas capsulata 28 D16147 DSM 30196T - Sphingomonas chlorophenolica X87161 DSM 7098T - Sphingomonas chlorophenolica RA2 X87164 DSM 6824 - Sphingomonas macrogoltabida 203 D13723 DSM 8826T - Sphingomonas mali Y-347 Y09638 DSM 10565T - Sphingomonas notatoria UQM2507 AB024288 DSM 3183T - Sphingomonas parapaucimobilis OH3607 D13724 DSM 7463T - Sphingomonas paucimobilis CL1/70 D13725 DSM 1098T - Sphingomonas rosa R135 D13945 DSM 7285T - Sphingomonas sanguis KM2397 D13726 LMG 2240T - Sphingomonas subartica KF1 X94102 DSM 10700T - Sphingomonas suberifaciens CR-CA1 D13737 DSM 7465T - Sphingomonas terrae E-1-A D13727 DSM 8831T - Sphingomonas trueperi X97776 DSM 7225T - Sphingomonas ursincola KR-99 AB024289 DSM 9006T - Sphingomonas wittichii RW1 AB021492 DSM 6014T - Sphingomonas xenophaga BN6 X94098 DSM 6383T - Sphingomonas yanoikuyae AB1105 D16145 DSM 7462T - Sphingomonas yanoikuyae B1 X94099 DSM 6900 - Sphingomonas yanoikuyae Pn4S D13946 LMG 3925 - Sphingomonas sp. EPA505 U37341 DSM 7526 + Sphingomonas sp. HV3 Y12803 (Yrjala et al. 1998) - (Bastiaens et al. 2000; - Sphingomonas sp. LB126 AF335501 Bastiaens et al. 2001) Sphingomonas sp. LH128 AY151394 (Bastiaens 1998) - Sphingomonas sp. LH227 AY151393 (Bastiaens et al. 2000) - Sphingomonas sp. VM0440 AY151392 (Springael et al. unpublished) - Sphingomonas sp. VM0506 AF335468 (Springael et al. unpublished) - Other Sphingomonadaceae genera Porphyrobacter neustonensis AB033327 DSM 9434T - Porphyrobacter tepidarius OT3 AB033328 DSM 10594T - Erythrobacter litoralis T4 AB013354 DSM 8509T - Erythromicrobium ramosum E5 AB013355 DSM 8510T - Zymomonas mobilis subsp. paniaceae I AF281032 LMG448T - Other α–Proteobacteria Agrobacterium luteum A61 NR DSM 5889T - Rhizobium rubi TR3 D12787 DSM 6772T - Brevundimonas diminuta 342 AJ227778 DSM 7234T - T = species type strain and/or genus type species. † result of PCR with specific primers EPAf and EPAr on soil DNA extract: + = PCR product, - = no detectable PCR product.

Soils used in this study. The soil samples used in this study were derived from different anthropogenic PAH-contaminated sites (Table 5-2). Chemical properties and analysis of the soil samples were described elsewhere (Chapter 2).

- 120 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils.

TABLE 5-2 SOIL SAMPLES USED IN THIS STUDY

Soil TOC PAH conc. Oil conc. DNA conc.* SPHINGO EPA Soil Origin pH -1 -1 -1 † † type (%) (mg kg ) (mg kg ) (µg g ) PCR PCR K3840 Gasoline station site (Denmark) Sand 8.2 0.50 20 98 2.75 +, (106) +, (106) B101 Coal gasification plant (Belgium) Sand 7.0 2.63 107 70 27.25 +, (105) +, (104) TM Coal gasification plant (Belgium) Sand 8.0 3.85 506 4600 4.75 +, (106) +, (105) Barl Coal gasification plant (Germany) Gravel 8.9 4.63 1029 109 6.15 +, (106) NP, (<104) AndE Railway station site (Spain) Clay 8.1 2.35 3022 2700 3.40 +, (106) +, (106) * DNA recovery per g of soil, mean value of 2 parallel extractions of 1 soil sample. † Result of PCR with genus specific primers Sphingo108f and Sphingo420r on soil DNA extract (Chapter 4). ‡ Result of PCR with species specific primers EPAf and EPAr on soil DNA extract: +, PCR product; NP, no detectable PCR product. Roughly estimated Sphingomonas cell concentration based on a ‘dilution to extinction’ PCR approach are indicated between brackets.

Analysis of phylogenetic position of Sphingomonas sp. strain EPA505. A multiple alignment of Sphingomonas 16S rRNA gene sequences was constructed using the Bionumerics software (Version 2.50, Applied Maths, Kortrijk, Belgium). The alignment consisted of approximately 275 sequences of both environmental and clinical Sphingomonas species available from the GenBank database (NCBI). Sequences were edited manually to remove nucleotide positions of ambiguous alignment and gaps. Sequence similarities were calculated, corrected using Kimura's two-parameter algorithm to compensate for multiple nucleotide exchange and used to construct a distance-based evolutionary tree with the Neighbor-Joining algorithm (Saitou et al. 1987). The topography of the branching order within the dendrogram was evaluated by using the Maximum-Likelihood and the Maximum-Parsimony character-based algorithms in parallel combined with bootstrap analysis with a round of 500 reassemblings. An out-group of the closely related α-Proteobacteria genera Rhizobium and Rhodospirillum were included to root the tree. Design of a new Sphingomonas sp. EPA505 specific 16S rRNA gene primer set. The specific primer set was selected in highly variable regions in the Sphingomonas 16S rRNA gene sequences. The forward primer EPAf (5’ CGAACGATCTCTTCGGAG 3’, E.coli position bp 62-80) and reverse primer EPAr (5’ TCAACAATCGTCCAGTGA 3’, E.coli position bp 758-776) were designed to amplify specifically a 696 bp fragment of the 16S rRNA gene. The theoretical selectivity of the EPA- primer pair was evaluated by visual analysis of the primer region within the constructed alignment of Sphingomonas rrn genes, by the Sequence Match program (RDP II) (Cole et al. 2003) and by the Advanced Blast Search program (Genbank, NCBI) (Altschul et al. 1990). DNA-extraction. Genomic DNA extraction from pure bacterial cultures and soil samples was performed as described previously (Chapter 2). PCR amplification of pure strain and soil DNA. To assure that the soil DNA was of good quality for PCR, dilution series of all soil DNA extracts were tested in PCR with universal eubacterial 16S rRNA gene primer pair GC40-63f and 518r as described previously (el Fantroussi et al. 1999). The PCR protocol using the EPAf and EPAr primer pair consisted of a short denaturation of 15 s at 95°C, followed by 50 cycles of denaturation for 3 s at 95°C, annealing for 10 s at 65°C and elongation for 30 s at 74°C. The last step included an extension for 2 min at 74°C. PCR was performed on Biometra (Whattman Biometra GmbH, Göttingen, Germany) or PerkinElmer (PerkinElmer, Wellesley, MA,

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USA) PCR-machines. PCR mixtures contained 100 ng of pure strain DNA or dilutions of soil DNA as templates, 1 U Taq polymerase, 25 pmol of the forward primer, 25 pmol of the reverse primer, 10 nmol of each dNTP, and 1 × PCR buffer in a final volume of 50 µl. Primers were synthesized by Westburg (Westburg BV, Leusden, The Netherlands). The Taq polymerase, dNTPs and PCR buffer were purchased from TaKaRa (TaKaRa Ex TaqTM, TaKaRa Shuzo Co., Ltd., Biomedical Group, Japan). Agarose gel electrophoresis of PCR products was performed using 1.5% agarose gels in 1x EY buffer with 50µg l-1 ethidiumbromide for 1 hour at 90V. Sensitivity of PCR detection. To examine the sensitivity of the PCR method to detect Sphingomonas sp. strain EPA505 in soil, a standard of living cells of Sphingomonas sp. strain EPA505 were added at different final cell concentrations (i.e. approximately 108, 106, 104, 102 CFU g-1) to PAH- contaminated soils (Table 2) prior to DNA-extraction. The total soil DNA extract was subsequently used as template in PCR with the EPAf and EPAr primers and PCR-products were analyzed by agarose gel electrophoresis. Detection of Sphingomonas sp. EPA505-related cells in soil. To roughly estimate the concentration of the detected Sphingomonas sp. EPA505 cells, dilution series of non-inoculated soil DNA extracts (1:1, 1:10, 1:100 and 1:1000 dilutions in water) were tested in a ‘dilution to extinction’ PCR-approach, similar to the MPN-PCR approach. The final cell concentration within a soil was deduced from the highest template dilution for which still a PCR product was detected, taking into account that the highest dilution giving a signal contained a cell density approaching the determined detection limit. In parallel, soil samples with inoculated Sphingomonas sp. EPA505 cells were regarded as positive PCR controls to assure that negative PCR results for samples without added cells were not due to PCR inhibition effects. Sequence analysis of amplified 16S rRNA gene fragments. PCR products derived soil DNA extracts with the specific primer set EPAf and EPAr were cloned into plasmid vector pCR2.1-TOPO using the TOPO Cloning Kit (N.V. Invitrogen SA, Merelbeke, Belgium) as described by the manufacturer. Clone inserts were tested again in PCR with the EPAf and EPAr primers and a 500 bp long fragment was sequenced by the Westburg Company (Westburg BV, Leusden, The Netherlands). A similarity analysis of the 16S rRNA gene sequences was done using the Blast Search program (Genbank, NCBI) (Altschul et al. 1990). Nucleotide sequence accession numbers. The partial 16S rRNA gene sequence of the Sphingomonas clones reported in this study are added to GenBank (NCBI) under accession numbers AY497362 to AY497377.

RESULTS

- 122 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils.

Phylogenetic position of Sphingomonas sp. strain EPA505. A phylogenetic tree for the Sphingomonas genus based on all currently available Sphingomonas 16S rRNA gene sequences was constructed (Figure 5-1). Twenty five of the 49 described PAH-degrading Sphingomonas strains were located in the branch of the former ‘Sphingobium’ genus (Takeuchi et al. 2001), i.e., the branch containing species S. yanoikuyae, S. clorophenolica, S. herbicidivorans, S. xenophaga, S. chungbukensis and S. cloaca. Based on 16S rRNA gene sequence, Sphingomonas sp. strain EPA505 (Mueller et al. 1990; Mueller et al. 1997), previously named Sphingomonas paucimobilis, was most related to PAH-degrading Sphingomonas sp. strains HS122 and HS163 (Han et al., unpublished) and Sphingomonas sp. strain G296-3 isolated from Antarctic ice (Christner et al. 2001). The closest related type T strain was the S. herbicidivorans DSM 11019 , which showed 96% 16S rRNA gene similarity with strain EPA505. Thus, strain EPA505 and its close relatives grouped in separated phylogenetic branch could possibly from a new species so far consisting only of PAH-degrading strains.

Design of a Sphingomonas sp. EPA505 specific 16S rRNA gene primer set. The EPAf and EPAr primer set was selected for PCR purposes. The primer set amplified a 696 bp fragment between bp 62 and bp 758 of the 16S rRNA gene according to the E. coli numbering. Blast (Genbank, NCBI) (Altschul et al. 1990) and Sequence Match (RDPII) (Cole et al. 2003) analysis showed that forward primer EPAf was 100% conserved in the 16S rRNA gene of Sphingomonas sp. strains EPA505 and in 5 additional 16S rRNA gene sequences of the 382 Sphingomonas present in the NCBI database (Figure 5-2). EPAf was 100% conserved in the 16S rRNA genes of Sphingomonas sp. strains G296-3, HS122, HS163 and non- Sphingomonas bacteria JP66.1 and K2-29 (Figure 5-2). Reverse primer EPAr was 100% conserved in the 16S rRNA gene of Sphingomonas sp. strain EPA505 and 13 other Sphingomonas gene sequences (Figure 5-2).

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12 10 8 6 4 2 0 AJ227778.1 .Brevundimonas diminuta LMG2089T

100 D30778.1 .Rhodospirillum rubrum ATCC11170T 96 AY345425.1 .Bacterium sp. K2-29 100 M11223.1 .Rhizobium radiobacter DMS30105 100 D14509.1 .Sinorhizobium meliloti IAM12611T AB033327.1 .Porphyrobacter nuestonensis DSM9434T 100 AB033328.1 .Porphyrobacter tepidarius DSM10594T 100 AB013355.1 .Erythromicrobium ramosum DSM8510T 99 M59062.1 .Erythrobacter longus DSM6997T 32 19 AB013354.1 .Erythrobacter litoralis ATCC700002T AB070237.1 .Sphingomonas tardaugens ATCC BAA-531T

77 AB025013.1 .Sphingomonas stygia CIP10514T 64 D16147.1 .Sphingomonas capsulata DSM30196T 82 AB025014.1 .Sphingomonas subterranea CIP105153T 99 100 AB025012.1 .Sphingomonas aromaticivorans DSM12444T * Y10678.1 .Sandaracinobacter sibiricus RB16-17 AF281032.1 .Zymomonas mobilis ATCC29192T AB024288.1 .Sphingomonas natatoria DSM3183T 100 AB024289.1 .Sphingomonas ursinicola DSM9006T

8 2 AY151394 Sphingomonas subartica LH128 * 100 X94102.1 .Sphingomonas subartica DSM10700T 5 D13727.1 .Sphingomonas terrae DSM8831T D13723.1 .Sphingomonas macrogoltabidus DSM8826T 99 64 Z73631.1 .Sphingomonas alaskensis DSM13593T 7 100 AB033949.1 .Sphingomonas witflariensis IFO15915 56 AF131297.1 .Sphingomonas taejonensis IFO16724T 79 AF367204.1 .Sphingomonas chilensis DSM14889T D84520.1 .Sphingomonas roseiflava CIP106847T 81 U37337.1 .Sphingomonas paucimobilis DSM10987T 100 D13724.1 .Sphingomonas parapaucimobilis DSM7463T 12 D13726.1 .Sphingomonas sanguinis ATCC51382T 2 60 AB021370.1 .Sphingomonas echinoides DSM1805T AB055863.1 .Sphingomonas melonisDSM14444T 100 AF131295.2 .Sphingomonas aquatilis IFO16772T 100 66 Y09639.1 .Sphingomonas asaccharolytica DSM10564T 99 Y09637.1 .Sphingomonas pruni DSM10566T 16 82 61 Y09638.1 .Sphingomonas mali DSM10565T D16146.1 .Sphingomonas adhaesiva DSM7418T 43 AF131296.1 .Sphingomonas koreensis IFO016723T D13945.1 .Sphingomonas rosa DSM7285T

10 AJ416411.1 .Sphingomonas hassiacum DSM14552T 32 AB021492.2 .Sphingomonas wittichii DSM6014T 75 6 X97776.1 .Sphingomonas trueperi DSM7225T 100 AJ243751.1 .Sphingomonas pituitosa DSM13101T D13737.1 .Sphingomonas suberifaciens DSM7465T 38 AY190158.1 .Sphingomonas sp. S32* 13 P@Sphingo. Sphingomonas sp. S34 U37345.1 .Sphingomonas sp. UN1F1* AF184222.1 .Sphingomonas sp. Ant17* 30 73 U37346.1 .Sphingomonas sp. UN1F2* 49 X94098.1 .Sphingomonas xenophaga DSM6383T* 100 35 AB025279.1 .Sphingomonas sp. Po626B4-1*

5 AY212326.1 .Sphingomonas sp. S21*

27 AY345575.1 .Uncultured bacterium sp. clone GLB-5 42 AY190157.1 .Sphingomonas sp. S7* 55 AY190138.1 .Sphingomonas sp. S10* AY151393 Sphingomonas sp. LH227*

35 1 AY190152.1 .Sphingomonas sp. S9* 96 90 AY190164.1 .Sphingomonas sp. S38.2* AY190134.1 .Sphingomonas sp. S2* 25 D16145.1 .Sphingomonas yanoikuyae DSM7462T

97 D13946.1 .Sphingomonas yanoikuyae LM G3925 78 X85023.1 .Sphingomonas yanoikuyae* 93 X94099.1 .Sphingomonas yanoikuyae B1* 2 U52146.1 .Sphingomonas sp. CF06* 29 AB040739.1 .Sphingomonas cloacae CIP107076T AB025530.1 .Sphingomonas sp. Lo251* 93 AB025572.1 .Sphingomonas sp. Lo41* 2 AY026948.1 .Sphingomonas sp. ML1* AY367016.1 .Sphingomonas sp. Sg6* Former 31 63 AY367015.1 .Sphingomonas sp. Sg5* 1 84 ‘Sphingobium’ 99 AY367017.1 .Sphingomonas sp. 12H6* cluster AF494538.1 .Sphingomonas sp. BPC4* U37341.1 .Sphingomonas sp. EPA505* (DSM7526) 5 68 AF395036.1 .Sphingomonas sp. G296-3 70 AY116880.1 .Sphingomonas sp. HS163* 98 AY116879.1 .Sphingomonas sp. HS122* AY460123.1 .Sphingomonas sp. ZL5 84 16 AF025350.1 .Sphingomonas sp. BRW 2°

42 AF191021.1 Sphingomonas sp. TA° 84 AF191022.1 .Sphingomonas sp. CD° 36 AB042233.1 .Sphingomonas herbicidovorans (DSM11019T)° 37 33 AJ252706.1 .Uncultured bacterium sp. clone RC-III-4 X87163.1 .Sphingomonas chlorophenolica (ATCC39723) 100 X87164.1 .Sphingomonas chlorophenolica RA2 (DSM6965)

67 X87162.1 .Sphingomonas chlorophenolica SR3 98 X87161.1 .Sphingomonas chlorophenolica (DSM7098T) 59 Y12803.1 .Sphingomonas sp. HV3*

65 AF159257.1 .Sphingomonas chungbukensis DJ77* (JCM11454T)

100 AF335468 Sphingomonas sp. VM 0506* 53 AF335501.1 .Sphingomonas sp. LB126* 92 AY151392 Sphingomonas sp. VM 0440*

FIGURE 5-1 POSITION OF PAH-DEGRADING SPHINGOMONAS SP. EPA505 IN THE 16S RRNA GENE BASED PHYLOGENETIC TREE

- 124 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils.

The evolutionary tree was generated by the Neighbor-joining method based on Kimura 2-parameter corrected similarity percentages and branching orders were evaluated using the Maximum-Parsimony algorithm. The topology was evaluated by bootstrap analysis (500 reassemblings) and percentages of bootstrap support are indicated at the branch points, with values above 70% indicating reliable branches. An out-group of the closely related α-Proteobacteria genera Rhizobium and Rhodospirillum were included to root the tree. The bar at the top indicates the estimated evolutionary distance, i.e., 1% indicating an average of 1 nucleotide substitution at any nucleotide position per 100 nucleotide positions. The evolutionary distance between two strains is the sum of the branch lengths between them. The former ‘Sphingobium cluster’ is indicated as well as the 6 Sphingomonas species type strains in this branch (arrows). PAH-degrading strains are indicated with * and pesticide degrading strains are indicated with °. The cluster of Sphingomonas sp. EPA505 related strains is highlighted in a box.

EPAf EPAr 5’ CGAACGATCTCTTCGGAG 3’ 5’... TCAACAATCGTCCAGTGA…3’

Sphingomonas sp. EPA505 (U37341) ------Sphingomonas sp. G296-3 (AF395036) ------Sphingomonas sp. HS122 (AY116879) ------No sequence available Sphingomonas sp. HS163 (AY116880) ------No sequence available Sphingomonas sp. Sg-5 (AY367015) No sequence available ------Sphingomonas sp. Sg-6 (AY367016) No sequence available ------Sphingomonas sp. 12H6 (AY367017) No sequence available ------Sphingomonas sp. TA (AF191021) No sequence available ------Sphingomonas sp. ZL5 (AY460123) No sequence available ------Bacterium clone RC-III-4 (AJ252706) No sequence available ------Sphingomonas clone SB46 (AY193926) No sequence available ------Sphingomonas clone GLB-5 (AY345575) No sequence available ------Bacterium JP66.1 (AY007677) ------C--TT---C ------TT--CGGC ----- Bacterium K2-29 (AY345425) ------T-----A-- GTT--AG ---- S. herbicidivorans DSM11019T (AB022428) ------GA------T------Sphingomonas sp. CD (AF191022) ------GA------T------Sphingomonas sp. BRW2 (AF025350) ------GA------T------S. cholorphenolica DSM7098T (X87161) ------GAC------GT------T - C-T - CA------G----- S. chungbukensis JCM11454T (AF159257) ------CT------G------T - C-T - C -T------C------S. xenophaga DSM6383T (X94098) ------GAC------GT------T - C-T - C -T------C------S. yanoikuyae DSM7462T (D16145) ------GA------T------T - C-T - CCA------S. cloaca JCM10874T (AB040739) ------GA------T------TGC-TGCCG------

FIGURE 5-2 DNA-SEQUENCE HOMOLOGY BETWEEN THE EPAF AND EPAR PRIMER SET AND THE 16S RRNA GENE SEQUENCES FROM DIFFERENT SPHINGOMONAS STRAINS Results are presented in a consensus table of matches. Dashes indicate homologous sequences.

Most strains with sequence homology to one of the two primers are phylogenetically very closely related to Sphingomonas sp. strain EPA505 and most of them are capable of degrading pesticides (strains TA, CD, BRW2) or polycyclic aromatic carbons (strains HS122, HS163, Sg-5, Sg-6, 12H6, BCP4) (Figure 5-1). Based on sequence homology, the primer set was thus expected to amplify only a fragment of the 16S rRNA genes from Sphingomonas sp. strains EPA505 and G296-3, and possibly some closely related strains of the EPA505 phylogenetic cluster for which the complementary sequences were unfortunately not available. The EPAf and EPAr

- 125 - Chapter 5 primer pair was tested on DNA from related and less related strains (Table 5-1). A PCR fragment was only obtained when using template DNA from strain EPA505 and not from the other tested Sphingomonas and non-Sphingomonas bacteria as was predicted. Sphingomonas sp. strains G296-3 and the other closely related strains with homologues 16S rRNA gene primer regions were unfortunately not available to be tested.

Detection limits of PCR protocol with primers EPAf and EPAr. The sensitivity of the new primer set EPAr and EPAr for detection of strain EPA505 was tested by adding viable cells of Sphingomonas sp. EPA505 at different final cell concentrations to several contaminated soils prior to DNA-extraction. Sphingomonas sp. strain EPA505 could be detected to a cell concentration of 104 and sometimes even 102 added cells per gram of soil.

Presence of Sphingomonas sp. EPA505 related strains in PAH-contaminated soils. The EPAf and EPAr primer set was used to examine 5 PAH-contaminated soil samples with different contamination records (Table 5-2) for the presence of Sphingomonas strains related to strain EPA505. A previous study had shown that Sphingomonas were present in all 5 tested soils (Chapter 4). Indigenous Sphingomonas sp. EPA505 related cells were detected in relatively high cell concentrations in 4 of the 5 tested PAH-contaminated soils (Table 5-2). Sequencing of 16 randomly chosen cloned PCR products obtained with the EPA primer set from soils K3840, B101 and AndE, confirmed that the detected signals were 16S rRNA gene fragments with 99% to 100% similarity to the 16S rRNA gene of Sphingomonas sp. EPA505.

DISCUSSION

- 126 - Occurrence of Sphingomonas sp. EPA505 in PAH-contaminated soils.

The Sphingomonas genus is a very interesting genus as it groups many bacterial isolates specialized in degradation of a wide variety of environmental pollutants. Sphingomonas sp. EPA505 is a model strain for the PAH-degrading Sphingomonas strains. Based on 16S rRNA gene sequence analysis Sphingomonas sp. EPA505 forms a separated branch in the phylogenetic tree of the Sphingomonas genus together with some other Sphingomonas strains, and may therefore represent a new Sphingomonas species. This should be confirmed by additional experiments such as FAME-analysis and total genome DNA-DNA hybridization. Mohn et al. (Mohn et al. 1999) concluded in 1999 that the phylogeny of the Sphingomonas genus was independent of the catabolic capabilities of its members, i.e., degradation of abietane triterpenoids. In our study, however, we found that all isolates closely related to the PAH-degrading strain Sphingomonas sp. EPA505, obtained from diverse PAH-contaminated soils from all over the world, have the capacity to degrade PAHs. For the culture-independent detection of Sphingomonas sp. EPA related strains, a new 16S rRNA gene based primer set, i.e., EPAf and EPAr primers, was developed. The new primer set was shown to be very specific for the target strain in PCR on pure strain and soil DNA extracts and could detect cell concentrations as low as 104 to 102 CFU g-1 soil. As such, the EPAf and EPAr primer set is the first specific Sphingomonas primer pair for the detection of a group of closely related PAH- degrading Sphingomonas sp. strains. The detection of Sphingomonas sp. EPA505 related cells in 4 out of the 5 tested PAH- contaminated soils clearly indicates the ubiquitous presence of this type of Sphingomonas strains in PAH-polluted soils. EPA505 related cells were detected in soils with both low and high concentrations of PAHs (100 – 3000 mg kg-1 soil). Cell concentrations were estimated between 104 and 106 cells per gram soil. Previously, via a PCR-DGGE method using a primer set targeting all members of the Sphingomonas genus, Sphingomonas sp. bacteria were detected in all 5 soils (Table 5- 2). Soils containing high PAH-concentrations showed less heterogeneous DGGE community fingerprints as the Sphingomonas populations in these soils were probably dominated by high numbers of a few Sphingomonas species. However, 16S rRNA gene sequences related to that of EPA505 were never found among the clone sequences retrieved from these soils by means of the general Sphingomonas specific primer set (Chapter 4). Moreover, EPA505-related cells were also never recovered from the same soils using a culture-based selective plating method (Vanbroekhoven et

- 127 - Chapter 5 al. unpublished). Detection methods targeting a larger group of Sphingomonas species are biased in favor of the most abundant species. A strain specific PCR-based detection method is thus really valuable and can reveal additional information on the Sphingomonas populations, detecting species that were otherwise missed. This study indicates that EPA505 related strains are certainly an important group of PAH-degrading Sphingomonas well adapted to harsh environmental conditions, and that they may have an important role in natural biorestauration of PAH-contaminated sites. In addition, such specialized bacteria can also be very useful for soil bioremediation by bioaugmentation. The newly developed strain specific detection method is thus indispensable to further study and monitor the presence and survival of this specific group of PAH-degrading Sphingomonas sp. EPA505 related cells in their environmental habitats. The new primers could also be useful for RT-PCR targeting rRNA to study more in detail the activity of the Sphingomonas sp. EPA505 related strains during PAH-biodegradation processes. Furthermore, the EPA primer set could be complementary to mRNA primers detecting the expression of Sphingomonas PAH catabolic genes in soil, to link degradation activity to the presence of certain Sphingomonas species (Story et al. 2000; Story et al. 2001; Pinyakong et al. 2003a). This will add to a better understanding of the role of Sphingomonas sp. EPA505 related strains in the biodegradation of PAHs in the environment.

- 128 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

CHAPTER 6

INFLUENCE OF THE CARBON/NITROGEN/PHOSPHATE-RATIO ON PAH-DEGRADATION BY MYCOBACTERIUM AND SPHINGOMONAS STRAINS IN SOIL

* REDRAFTED AFTER: LEYS NATALIE, BASTIAENS LEEN, VERSTRAETE WILLY, SPRINGAEL DIRK

(SUBMITTED) INFLUENCE OF THE CARBON/NITROGEN/PHOSPHATE-RATIO ON PAH-DEGRADATION BY

MYCOBACTERIUM AND SPHINGOMONAS STRAINS IN SOIL, APPL. MICROBIOL. BIOTECHNOL.

ABSTRACT

Biodegradation of polycyclic aromatic hydrocarbons (PAHs) in the environment is still often limited due to unfavorable nutrient conditions for the bacteria that use these PAHs as sole source of carbon and energy. Mycobacterium and Sphingomonas are 2 PAH-degrading specialists common present in PAH-polluted soil, but not much is known about their specific nutrient requirements. By adding different inorganic supplements of nitrogen (N) and phosphorus (P) affecting the overall Carbon/Nitrogen/Phosphorus-ratio of soil in soil slurry degradation tests, we investigated the impact of soil inorganic N and P nutrient conditions on PAH degradation by PAH-degrading Sphingomonas and Mycobacterium strains. The general theoretical calculated C/N/P-ratio of 100/10/1 [expressed in mole] allowed rapid PAH metabolisation by Sphingomonas and Mycobacterium strains without limitation. In addition, PAH-degradation activity was not affected when circa 10 times lower concentrations of nitrogen and phosphorus were available, indicating that Sphingomonas and Mycobacterium strains are capable of metabolizing PAHs under low nutrient conditions. Nor does PAH-degradation seemed to be affected by excesses of nitrogen and phosphorus unbalancing the C/N/P ratio in the soil. Supplements of nitrogen and phosphorus salts increased however the salinity of soil slurry solutions and seriously limited or even completely blocked biodegradation.

- 129 - Chapter 6

INTRODUCTION

Polycyclic aromatic hydrocarbons (PAHs) are common pollutants of contaminated industrial sites found in industrialized countries (Edwards 1983). PAHs are of environmental concern due to their toxic, mutagenic and carcinogenic properties (Enzminger 1987). Microbial degradation is considered as a major route through which PAHs are naturally removed from contaminated environments (Bossert et al. 1984; Cerniglia 1984; Enzminger 1987; Cerniglia 1992). Therefore, bioremediation is considered as an economically and ecologically beneficial remediation technology for the treatment of PAH polluted sites (Erickson et al. 1993; Wilson et al. 1993; Luthy et al. 1994; Würdemann et al. 1995). Although, PAH-degrading microorganisms are often naturally present in PAH- polluted soil (Cerniglia 1984; Cerniglia 1992; Juhasz et al. 2000b; Kanaly et al. 2000b), the rate of PAH-biodegradation in the field is mostly low in comparison to degradation rates obtained in laboratory culture conditions (Erickson et al. 1993; Luthy et al. 1994). Many different soil parameters such as oxygen concentration, redox potential, moisture, acidity, salinity, temperature, mineralogy and granularity, contamination profile and competition for nutrients between the indigenous microflora could constrain the PAH biodegradation in the natural soil environment (Wilson et al. 1993; Beck et al. 1995). One of the most important factors may be the limited availability of nutrients in the soil. The principle in PAH-bioremediation is that heterotrophic soil microorganisms will use the carbon supplied by the PAHs for growth and energy. However, microorganisms depend besides on organic compounds as carbon sources (C) also on inorganic macronutrients like nitrogen (N) and phosphor (P) for cell growth and energy production. In the field, mostly the C/N/P- conditions are far from optimal. The argument that competition for nitrogen by plant roots, mycorhizal fungi and microorganisms keep inorganic N and P concentrations low is widely accepted. Engineers and microbiologists have tried to manipulate the soil system to increase biodegradation rates. Addition of nutrients (biostimulation) and/or bacteria (bioaugmentation) is the most common used strategy but with variable and unpredictable success. Addition of nitrogen and phosphate compounds has been practiced to balance insufficient inorganic nutrient concentrations with the CPAH- concentrations in soil. Addition of inorganic nutrients has been reported to both

- 130 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas. positively affect and negatively effect different aspects of pollutant degradation kinetics such as lag time, degradation rate and degradation extent (Jobson et al. 1974; Bossert et al. 1984; Thorton-Manning et al. 1987; Swindoll et al. 1988; Johnson et al. 1990; Manilal et al. 1991; Morgan et al. 1992; Alexander 1994; Baker et al. 1994;

Carmichael et al. 1997; Ward et al. 1999). Many biostimulation and bioaugmentation degradation tests were succesfull on well-controlled lab scale and even on pilot scale

(Wilson et al. 1993; Cookson 1995; Bastiaens 1998) but often failed on a larger field scale (Gemoets et al. 2000). In addition, inorganic supplements of nitrogen and phosphorus can be very effective in closed systems (lab scale and pilot scale) but have the tendency to wash out in outdoor field applications (Leahy et al. 1990). All these studies indicate that PAH-biodegradation in soil is still very much a ‘black box’ system. Not much is known about the specific needs for groups of bacteria specialized in PAH degradation for optimal activity in the soil environment. A more general approach is needed to reveal general conditions at which certain groups of PAH-degrading bacteria work best in soil. Mycobacterium and Sphingomonas strains have been often isolated as PAH-degraders from PAH-contaminated soil (Bastiaens et al. 2000) and have been identified as major colonizers of such soils (Chapter 2, 3, 4, 5) indicating that they may be important sinks of PAHs in the soil environment. It is thus of interest to optimize the growth and activity of these two genera. Therefore, we investigated in detail the influence of different nutrient conditions on the survival and activity of PAH-degrading Mycobacterium and Sphingomonas strains in different soils. Different PAH-degrading Mycobacterium and Sphingomonas strains, previously isolated from historically contaminated soils, were used as model strains in this research. Degradation tests were set up in small-scale microcosms simulating soil slurry reactor conditions, to examine the response of seeded (and indigenous) Mycobacterium and Sphingomonas strains to different mineral inorganic nutrient supplements in soil.

- 131 - Chapter 6

MATERIALS AND METHODS

Bacterial strains and cultivation conditions. Three different Sphingomonas strains and three different Mycobacterium strains were used previously selected for their ability to completely mineralize different PAHs (Bastiaens 1998; Bastiaens et al. 2000; Springael et al. unpublished). Sphingomonas sp. LH128, LB126 and EPA505 grow on respectively phenanthrene, fluorene and fluoranthene, while Mycobacterium sp. LB307T, LB501T and VM552 grow respectively on phenanthrene, anthracene and pyrene (Table 6-1). Inoculum strains were precultured on 2 g l-1 PAH crystals as only carbon source in liquid mineral medium (Wick et al. 2001). Cells for inoculation were harvested from cultures in stationary phase. Prior to inoculation the remaining PAH-crystals were filtered out over a glass wool filter and cells were washed 2 times in an aqueous solution of 0.85% NaCl.

TABLE 6-1 BACTERIAL STRAINS USED IN THIS STUDY AND THEIR PAH-DEGRADATION CHARACTERISTICS

Strain Grows on References Sphingomonas sp. LB126 Fluorene (Bastiaens 1998; Bastiaens et al. 2000; Bastiaens et al. 2001; Wattiau et al. 2001; van Herwijnen et al. 2003b) Sphingomonas sp. LH128 Phenanthrene (Bastiaens 1998; Bastiaens et al. 2000) Sphingomonas sp. EPA505 Fluoranthene (Mueller et al. 1990; Story et al. 2000; Story et al. 2001) Mycobacterium sp. LB307T Phenanthrene (Bastiaens 1998; Bastiaens et al. 2000; Willumsen et al. 2001b) Mycobacterium sp. LB501T Anthracene (Bastiaens 1998; Bastiaens et al. 2000; Wick et al. 2001; Wick et al. 2002a; Wick et al. 2002b; van Herwijnen et al. 2003a; Wick et al. 2003b) Mycobacterium sp. VM0552 Pyrene (Wick et al. 2002b; Springael et al. unpublished)

Soil samples. The analytical characteristics of the soil samples used in this study are reported in Table 6-2. PAH-contaminated soil samples B101 and K3840 were obtained from different historically PAH- contaminated industrial sites and analyzed as described previously (Chapter 2). In addition, one non- sterile well-characterized non-contaminated sandy soil (MOL) was used originating from a nature reserve in Mol, Belgium, lacking a natural PAH degrading potential. It contained a normal natural + - amount of inorganic available nitrogen (N) (N-NH4 and N-NO3 ) and phosphorus (P) and a relatively high natural sulfur (S) concentration.

TABLE 6-2 PHYSICO-CHEMICAL CHARACTERISTICS OF THE SOILS USED IN THIS STUDY.

Parameter MOL B101 K3840 Water content % 3.62 16.50 6.71 pH 6.80 7.00 8.30 Carbon % TC 1.21 2.63 1.07 % TOC 1.20 2.63 0.83 % TIC 0.01 < 0.01 0.24 Nitrogen mg kg-1 DW total N 987 1270 334 -1 + mg l N-NH4 (1/10 soil/water) <1.00 <1.00 <1.00 -1 2- mg l NO3 (1/10 soil/water) 21.20 5.18 7.42 -1 3- Phosphor mg l PO4 (1/10 soil/water) 2.14 13.00 6.28 -1 2- Sulphur mg l SO4 (1/10 soil/water) 40.20 1.59 8.55 Metals mg kg-1 DW Fe3+ 81600 12100 93600 mg kg-1 DW Zn2+ 31 155 62 Contaminants mg kg-1 DW cyanide 0.60 1.96 <0.25 mg kg-1 DW mineral oil <50 950 80 mg kg-1 DW 17 EPA-PAHs <0.01 108 66

Natural C/N/P ratio CPAH/N/P [mg/kg] 0/48/7 108/12/42 66/17/21 C/N/P ratio after spiking with CPAH/N/P [mg] 120/10/1 120/2/9 120/3/4 -1 600mg kg PAH CPAH/N/P [mole] 100/7/0.5 100/2/3 100/2/1 TC = Total Carbon, TOC = Total Organic Carbon, TIC = Total Inorganic Carbon, DW = dry weight

- 132 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

Experimental set up of small scale soil slurry PAH biodegradation experiments. For the biodegradation tests, small-scale shaken soil slurry systems were used as previously described

(Bastiaens 1998). Glass tubes (10ml) with aluminum-lined screw caps contained 0.5 gram of soil. The soil was artificially contaminated with 600 mg kg-1 of a single PAH by spiking small volumes (30 to 60 µl) of concentrated acetone-PAHs solutions directly on the dry soil matrix using glass syringes and by evaporating the acetone overnight. A volume of 2.5 ml aqueous solution was added to create slurry with 17% dry weight. The composition of this aqueous solution depended on the additives that were tested. The soil slurry systems were inoculated with 0.5ml of a cell suspension with optical density of around 0.300. The 3 ml soil slurries, containing approximately 0.3 mg of a single PAH (approximately 0.283 mg C) with approximately108 cells of a single PAH-degrading bacterial strain, were incubated at 30°C while shaking (200 rpm). The degradation tests were done in a ‘batch set up’, i.e., a series of identical reaction vials for each test condition were set up. For each analysis point in time, one vial of a series of identical vials was used to measure the residual PAH-concentration, pH and ionic strength (I). Changing the nutrient conditions in the soil slurry. The Carbon/Nitrogen/Phosphor ratio of the soil slurry was modified, by the addition of NH4Cl as nitrogen source and K2HPO4 as phosphor source in accordance to the PAH-concentration as C-source. The ionic strength of the soil solution was modified by the addition of MgCl2. In a positive control series for maximal stimulation of the degradation activity, a phosphate buffered mineral growth medium containing spore elements was provided (MM) (Wick et al. 2003b). In addition, two negative control series were included in each degradation experiment, one without inoculated cells and one containing formaldehyde and sodium azide metabolically inactivated cells to check and correct for abiotic removal due to sorption, oxidation, etc. All amendment solutions were adjusted to the same neutral pH of the phosphate buffered medium (MM), i.e., a pH of 7.4, to exclude the interference of pH–effects. The ionic strength of the solutions differed. Other parameters were kept optimal, i.e., all vials were incubated at optimal temperature (30°C) and oxygen supply (orbital shaken at 200rpm). Monitoring biodegradation activity. To monitor the biodegradation kinetics of the inoculum strain under the pre-set conditions, the residual PAH concentration was measured at different times after inoculation. PAHs were 2 times extracted with 3ml hexane and the extracts were pooled and analyzed by HPLC (Merck Hitachi LACHROM pump L-7100). Of each sample, 20µl was automatically injected (Marathon autosampler) on a 5µm LiChrospher 100 RP-18 encapped column and PAHs were eluted with a mixture of 25% water and 75% acetonnitril at a flow rate of 1ml per minute. The PAHs in the eluent were detected at 254nm using a UV-VIS detector (Merck Hitachi L-4250). For the integration of the chromatograms and quantification of the PAH-concentration, the software packet BORWIN (ATAS) was used. Degradation kinetics were plotted as the disappearance of the PAH-compound as a function of time. At each sampling point in time two vials were sacrificed for complete analysis, so that each analysis result represented duplicates. Monitoring acidity (pH) and salinity (I). During biodegradation tests the pH and I of each test series was monitored in a parallel vial incubated under identical test conditions. Measurements were done by the immersion of a pH-electrode into the aqueous solution of the soil slurry microcosms after the soil

- 133 - Chapter 6

was let to sediment for 5 min. The measurement was registered by a pH meter. The salinity or ionic strength (I) of the soil suspensions was recorded as electrical conductivity (µS/cm) using a standard conductivity meter. The I was calculated using the equation: I = 1.6x10-5x conductivity (µS/cm). Monitoring bacterial growth. Cell multiplication of the inoculated bacteria in the soil slurry was followed by plating the soil slurry supernatant on solid medium. Some test vials were used for total DNA extraction and PCR with 16S rDNA specific primers.

RESULTS

PAH mineralization by Mycobacterium and Sphingomonas strains in soil slurry under theoretically predicted optimal C/N/P-ratio. We evaluated practically whether or not the theoretically calculated optimal C/N/P ratio of 100/10/1 [mole] would also be optimal for growth of by Mycobacterium and Sphingomonas strains at the expense of PAHs in soil. By adding simple deionised -1 water to the PAH spiked (600 mg kg ) Mol soil, a CPAH/N/P-ratio of 120/10/1 [mg] or 100/7/0.5 [mole] was created based on the natural available N and P in the soil, approaching the theoretically predicted optimal ratio. This nutrient condition allowed indeed rapid and complete removal of fluorene by Sphingomonas sp. LB126, fluoranthene by Sphingomonas sp. EPA505, phenanthrene by Sphingomonas sp. LH128, anthracene by Mycobacterium sp. LB501T, phenanthrene by Mycobacterium sp. LB307T and pyrene by Mycobacterium sp. VM552 (Figure 6-1). The soil buffered + - the carbonate (H & HCO3 ) formation from the produced CO2 during bioremediation and suitable pH ranges between 6.0 and 7.0 were maintained (data not shown). Probably due to the difference in growth rate, PAH-degradation by Mycobacterium strains occurred generally at slower rates than degradation by Sphingomonas strains. The PAH-degradation rates could not be increased by adding the well-balanced MM medium to the soil (Figure 6-1) and degradation rates in soil slurry were identical to rates obtained in pure liquid tests in MM medium (data not shown). These results indicate that the degradation rates achieved at a C/N/P ratio of circa 120/14/3 [mg] or 100/10/1 [mole] in soil slurry were maximum.

- 134 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

Fluorene degradation by LB126 in soil slurry Fluoranthene degradation by EPA505 in soil slurry

120 120

100 100 ) ) %

0 80 o % C 80 / C / C ( (C c 60 max degradation rate 60 max degradation rate n = 1.6 mg/hour = 1.8 mg/hour o onc c c -

40 -

H 40 H A PA 20 P 20

0 0 0 24487296120144 0 24487296120144 Time (hours) Time (hour)

Phenanthrene degradation by LH128 in soil slurry Anthracene degradation by LB501T in soil slurry

120 120

100

) 100 ) % % o 80 0

C 80 C / / C ( (C . 60 . max degradation rate max degradation rate 60 = 0.9 mg/hour onc

= 3.7 mg/hour onc c c -

40 -

H 40 H A A P 20 P 20

0 0 0 24 48 72 96 120 144 0 48 96 144 192 240 288 336 Time (hour) Time (hour)

Pyrene degradataion by VM552 in soil slurry Phenanthrene degradatation by LB307T in soil slurry

120 120

100

) 100 ) 0 % 80 0 % 80 /C /C C max degradation rate C max degradation rate . ( . (

c = 1.3 mg/hour = 1.8 mg/hour 60 c 60 n n o co c - 40 - 40 H H A A P 20 P 20

0 0 0 24487296120144 0 24487296120 Time (hour) Time (hour)

inoculum + demin. w ater inoculum + MM dead inoculum + MM no inoculum + MM

PAH-conc = 600 mg/kg , Inoculum conc. = 2x108 cells/g

FIGURE 6-1 PAH-DEGRADATION BY SPHINGOMONAS SP. STRAINS LB126, EPA505 AND LH128 AND MYCOBACTERIUM SP. STRAINS LB501T, LB307T AND VM552 UNDER THEORETICALLY PREDICTED OPTIMAL C/N/P CONDITIONS IN MOL SOIL SLURRY

- 135 - Chapter 6

By adding different amounts of cells it was proven that the observed degradation was not only due to the addition of relative high cell concentrations with highly induced PAH-degradation enzymes but that this nutrient ratio also allowed easy production of new cells at the expense of the PAHs as source of carbon and energy (Figure 6-2). The addition of fewer cells, e.g. 10 cells per gram soil, of Mycobacterium sp. LB501T and VM0552 prolonged the lag phase for significant biodegradation of anthracene respectively pyrene under these conditions, but similar degradation rates were achieved after reaching similar cell densities. Monitoring of viable cell concentrations via selective plating on PAH-coverd minimal medium (data not shown), idicated that only initial or built up inoculum concentrations of circa 107 cells per gram soil could catalize significant degradation of the spiked PAHs. In addition, when the concentration of the pollutant was reduced in time, also the multiplication of the bacteria slowed down and cell concentrations finally declined (data not shown).

Antracene degradation by LB501T in soil slurry Pyrene degradation by VM552 in soil slurry

120 120 ) ) 100 100 % %

0 /C0 /C 80 80 C C . ( . ( c c n

n 60 60 co co - - H H 40 40 A A P P 20 20

0 0 0 48 96 144 192 240 288 336 0 48 96 144 192 240 288 336 384 Time (hour) Time (hour) PAH-conc = 600 mg/kg , Additive = distilled water

inoculum 6,0E+7 cells/g inoculum 6,0E+5 cells/g inoculum 6,0E+3 cells/g inoculum 6,0E+1 cells/g no inoculum dead inoculum

FIGURE 6-2 PAH-DEGRADATION BY MYCOBACTERIUM SP. STRAINS LB501T AND VM552 INITIATED WITH DIFFERENT INOCULUM CONCENTRATIONS IN MOL SOIL SLURRY

Liquid cultures in Wick medium showed that each 1 mg of carbon of PAHs (phenanthrene, fluorene respectively fluoranthene) could be converted in circa 106 cells of Sphingomonas sp. LH128, LB126 and EPA505. Thus, the circa 300 mg of PAHs added in the soil slurry degradation experiments could sustain a bacterial population of circa 3x108 cells or 1 generation of the added inoculum of 1.5x108

- 136 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

Sphingomonas cells. The growth rates of Mycobacterium sp. LB307T and LB501T on phenanthrene respectively anthracene in liquid cultures could not be measured by variation in optical density due to biofilm formation on the PAHs crystals as reported previously (Wick et al. 2001; Wick et al. 2002a).

Influence of natural unbalanced low nitrogen and phosphor concentrations on PAH-degradation by Mycobacterium spp. and Sphingomonas spp.

We calculated the natural CPAH/N/P-ratios for 10 industrial PAH-contaminated soil samples from different origin and containing different mineral and PAH concentrations (Table 6-3). In all analyzed soils, nitrogen concentrations were far below optimal concentration and in the 7 highest contaminated samples in addition phosphorus concentrations were low. The only soluble and thus bio-available form of inorganic nitrogen was nitrate except for one soil.

TABLE 6- 3 NATURAL C/N/P-RATIOS IN PAH-CONTAMINATED SOILS FROM DIFFERENT ORIGIN

Soil Origin CPAH/N/P CPAH/N/P CPAH/N/P IS [mg/kg] [mg] [mole] And E railway wood treatment site, Spain 3022/BDL/6 120/BDL/0.2 100/BDL/0.07 ND TB7 (30-70cm) coal gasification factory site, Belgium 2175/19/45 120/1/3 100/0.8/0.8 ND Barl coal gasification factory site, Germany 1109/1/BDL 120/0.1/BDL 100/0.09/BDL ND TB7 (0-30cm) coal gasification factory site, Belgium 650/19/0.7 120/4/0.1 100/3/0.04 ND TD coal gasification factory site, Belgium 516/3/BDL 120/0.8/BDL 100/0.5/BDL 0.002 And C railway wood treatment site, Spain 358/BDL/3 120/BDL/1 100/BDL/0.4 ND E6068 soil remediation company, Denmark 278/6/BDL 120/3/BDL 100/2/BDL ND B1 coal gasification factory site, Belgium 136/6/29 120/5/26 100/3/8 0.002 B101 coal gasification factory site, Belgium 108/12/42 120/13/47 100/9/15 0.001 TB3 coal gasification factory site, Belgium 14/BDL/1 120/BDL/10 100/BDL/3 ND OPTIMAL 120/14/3 100/10/1 -1 - - -1 + -1 3- 3- BLD = below detection limit: <1 mg l for NO3 (<2.3 mg N-NO3 ); <1 mg l N-NH4 ; <0.15 mg l for PO4 (<0.5 mg P-PO4 ) IS = Ionic Strength measured as conductivity, a indication of salinity ND = not determined

We investigated whether or not imbalances of low nitrogen and phosphor concentrations indeed still allowed fast PAH biodegradation by Sphingomonas and Mycobacterium spp in two different set ups. First, by adding high concentrations of -1 PAHs (1200 mg kg ) to the Mol soil normally rich in N and P, a CPAH/N/P-ratio of

120/3.4/0.5 [mg] or 100/2.4/0.7 [mole] with theoretically limiting nitrogen and phosphorus concentrations was created based on the natural available N and P in the soil. The nutrient condition in this set-up was clearly reducing PAH-degradation rates (Figure 6-3). The degradation of phenanthrene by Sphingomonas sp. LH128 and Mycobacterium sp. LB307T and of anthracene by Mycobacterium sp. LB501T under these conditions was much slower than in soil slurry enriched with a well balanced

- 137 - Chapter 6 minimal medium instead of with water. However, it seemed that other factors than the low N and P concentrations were causing the constraint since adding extra N and/or P could not stimulate degradation. Possibly, by adding large amounts of PAHs other parameters such as low spore element concentrations or soil buffer capacity could have become limiting or some toxic effects of high PAH-concentrations could start playing.

Phenanthrene degradation by LH128 in soil slurry Phenanthrene degradation by LB307T in soil slurry Spiked PAH-conc = 2100 mg/kg Spiked PAH-conc = 2100 mg/kg 120 120

100 100 ) ) % %

80 0 80 /C0 /C C C . ( . ( c

60 c 60 n n o c co 40 - 40 H AH- A P P max degradation rate 20 max degradation rate 20 = 2.2 mg/hour = 1.9 mg/hour 0 0 0 48 96 144 192 240 0 48 96 144 192 240

Time (hour) Time (hour)

Phenanthrene degradation by LH128 in soil slurry Antracene degradation by LB501T in soil slurry Spiked PAH-conc = 6000 mg/kg Spiked PAH-conc = 6000 mg/kg 140 120

) 120 max degradation rate 100 % ) = 2.0 mg/hour max degradation rate 0 = 2.5 mg/hour 100 % C 0 80 /C (C/ . 80 C c . ( n

c 60 o 60 n o c - 40 H AH-c 40 A P P 20 20

0 0 0 96 192 288 384 480 576 672 0 96 192 288 384 480 576 672 Time (hour) Time (hour)

inoculum + w ater inoculum + w ater + N inoculum + w ater + P inoculum + w ater + N & P inoculum + MM no inoculum + MM dead inoculum + MM

Inoculum concentration = circa 2x108 cells/g

FIGURE 6-3 PAH-DEGRADATION BY SPHINGOMONAS SP. STRAINS LH128 AND MYCOBACTERIUM SP. STRAINS LB501T AND LB307T UNDER THEORTICALLY LIMITING N/P CONDITIONS IN MOL SOIL SLURRY

- 138 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

Therefore, in a second test set up, moderated amounts of PAHs (600 mg kg-1) and inoculum cells were added to 2 different industrial PAH-contaminated soils naturally poor in N and P (soils K3840 respectively B101) creating ratio’s low in N (100/2/1 [mole] respectively 100/2/3 [mole]). Nitrogen depletion did not seem to limit fluorene degradation by Sphingomonas sp. LB126 at all in both tested soils (Figure 6-4). PAH- biodegradation was rapid and complete. Meanwhile low concentrations of other non- spiked PAHs in these industrial contaminated soils were not metabolized by the added strain (data not shown), possibly due to limited bioavailability. Both soils seemed to provide enough buffer capacity to maintain pH within optimal ranges during the degradation process (data not shown). The degradation rate of fluorene could not be stimulated by adding a well balanced mineral nutrient medium, indicating that the degradation rates achieved at low CPAH/N/P ratios were maximal.

Fluorene degradation by LB126 in K3840 soil slurry Fluorene degradation by LB126 in B101 soil slurry

120 120

100 100 ) ) % 0 0 % 80 80 /C /C C C . ( . (

c 60 c 60 n n o co c - 40 max degradation rate - 40 max degradation rate H = 0.5 mg/hour H = 0.4 mg/hour A P PA 20 20

0 0 0 48 96 144 192 240 04896144192240

Time (hour) Time (hour)

inoculum + w ater inoculum + MM no inoculum + w ater

no inoculum + MM dead inoculum + MM

Inoculum concentration = circa 2x108 cells/g

FIGURE 6-4 PAH-DEGRADATION BY SPHINGOMONAS SP. STRAIN LB126 UNDER THEORTICALLY LIMITING N/P CONDITIONS IN K3840 AND B101 SOIL SLURRY

- 139 - Chapter 6

Influence of imbalanced high nitrogen and phosphor concentrations on PAH- degradation by Mycobacterium and Sphingomonas strains in soil. We tested whether imbalanced high nitrogen and phosphor concentrations could have an effect on PAH biodegradation. By adding nitrogen as NH4Cl and/or phosphorus as

K2HPO4 to the PAH-spiked MOL soil, the CPAH/N/P-ratio was changed creating higher nitrogen and/or phosphor concentrations than would be optimal according to the 100/10/1 [mole] ratio. The internal N/P ratio remained constant at 10/1 [mole] or at 100/1 [mole]. The excess of nitrogen and phosphor according to the carbon or the excess of nitrogen according to phosphate and carbon concentrations did not have any positive nor negative effect on the fluorene or anthracene degradation kinetics of Sphingomonas sp. LB126 respectively Mycobacterium sp. LB501T in the soil slurry as long as not too much salt was added (Figure 6-5). Addition of a 1000 times more nitrate and phosphate than would be theoretically needed, completely blocked the fluorene, phenanthrene and fluoranthene degradation by Sphingomonas sp. LB126, LH128 and EPA505 and phenanthrene and anthracene degradation by Mycobacterium sp. LB307T and LB501T (Figure 6-5)(not all data shown). This was due to the increased salinity of the soil slurry (IS of 1.8) as demonstrated in an additional test were mineral medium was added with MgCl2 to bring the I of the soil slurry to the same I conditions as in the previous test with very high N- and P-concentrations (data not shown). In addition, liquid growth tests proved the inability of all test strains to grow in conditions with I equal or higher than 0.500, except for strain LB501T which was resistant to I of 0.500 but sensitive to I of 0.800 (data not shown). Strain LH128 was more sensitive than the other tested strains as it could not grow in medium with I of 0.250 (data not shown).

- 140 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas.

Fluorene degradation by LB126 in MOL soil slurry Anthracene degradation by LB501T in MOL soil slurry

120 120

100 100 ) ) %

% 80 0 80 /C /C0 C C

max degradation rate . ( max degradation rate . ( c c 60 = 0.7 mg/hour 60 = 0.2 mg/hour n n o co c 40 - 40 H A AH- P P 20 20

0 0 0 24487296120144 0 48 96 144 192 240 288 336 384 Time (hour) Time (hour)

inoculum + w ater (120/10/1.5) inoculum + w ater&N& P (120/22/2.7) inoculum + w ater&N&P (120/130/14) inoculum + w ater&N&P (120/1210/122) inoculum + w ater&N&P (120/12010/1202) inoculum + w ater&N&P (120/130/2.7) inoculum + w ater&N&P (120/1210/14) inoculumm + MM dead inoculum + MM no inoculum + MM

Inoculum conc. = circa 2x108 cells/g

FIGURE 6-5 PAH-DEGRADATION BY SPHINGOMONAS SP. STRAIN LB126 AND MYCOBACTERIUM PHINGOMONAS SP. STRAIN LB501T UNDER INCREASING N/P CONCENTRATIONS MOL SOIL SLURRY. The final theoretical CPAH /N/P-ratios obtained in the soil slurry, including the natural available N and P, are indicated between brackets.

DISCUSSION

As any other technology, engineered bioremediation of PAH-contaminated sites can only be successful if the active system is known and relative controllable, i.e., if the degrading soil microorganisms are identified, can be promoted in the creation and maintenance of an active biomass and can be directed towards optimal PAH- biodegradation. PAH-degrading Mycobacterium and Sphingomonas strains have repeatedly been isolated from and detected in PAH-contaminated soils (Bastiaens et al. 2000) (Chapter 2, 3, 4, 5) indicating that these bacteria may have an important role in the PAH-biodegrading soil microflora. In this study, we investigated the effect of inorganic nutrient availability on the degradation of PAHs in soil by Mycobacterium and Sphingomonas strains under relative optimal test conditions. The influences of temperature, humidity and PAH-

- 141 - Chapter 6 bioavailability on biodegradation were minimized as the PAHs were freshly added to the soil (no aging effects), the soil slurry was intensively mixed and incubated at 30°C during the whole degradation processes. It was demonstrated that the PAH- degradation by Sphingomonas and Mycobacterium strains in soil slurry was found maximal under carbon/nitrogen/phosphate concentrations of circa 120/14/3 [expressed in mg] or 100/10/1 [expressed in mole], i.e., a ratio theoretical predicted as optimal for cell growth in general and mostly used for biostimulation in soil bioremediation processes (Paul et al. 1989; Wilson et al. 1993; Hoeppel et al. 1994; Bouchez et al. 1995; Cookson 1995). Under these optimal nutrient conditions, metabolisation of PAHs sustained new biomass production by Sphingomonas and Mycobacterium cells. Mineralization curves of low inoculum concentrations were sigmoid in nature. As inoculum cells with fully activated PAH-catabolic enzyme systems were used, this indicates that a period of biomass production was needed for significant mineralization of the added PAH (Grosser et al. 1991; Macleod et al. 2002). Significant PAH removal was detectable when cell concentrations reached circa 107 cells per gram of soil. Similarly, p-nitrophenol disappearance in artificially contaminated sewage water was only detectable when cell concentration reached 105 CFU ml-1 (Wiggins et al. 1987). The addition of inorganic nutrients to balance soil C/N/P ratios is a general bioremediation practice. Our results indicate, however, that PAH-biodegradation by Sphingomonas and Mycobacterium spp. was not constrained by imbalanced

CPAH/N/P-ratios with low nitrogen and/or phosphate concentrations naturally occurring in most contaminated soils. It has been suggested that low inorganic nutrient concentrations in soil could reduce the number of active PAH-degrading bacteria to an insufficient level, since the slower growing pollutant degrading bacteria are out competed (Cerniglia 1992; Steffensen et al. 1995). Our results indicate, however, that the specific pollutant utilization and metabolisation capacities of PAH- degrading Sphingomonas and Mycobacterium strains might give them a selective advantage in polluted soils, making them competitive with faster growing bacteria. Our results could be consistent with the low overall C/N/P ratio of 120/2/0.15 [mg] or 100/1.3/0.05 [mole] that was suggested once as optimal for PAH-degradation in soil (Wilson et al. 1993). This would indicating that PAH-degrading bacteria would need 10 times less nitrogen and phosphorus because more PAH carbon is used for energy production instead of biomass production than normally for other substrates for which

- 142 - Influence of C/N/P-ratio on PAH-degradation by Mycobacterium and Sphingomonas. a ratio of 120/14/3 is suggested. This phenomenon has also been reported for bacterial growth on other xenobiotics. In addition, it is reported for Mycobacterium sp. LB501T that more CO2 is produced for cell maintenance and less for cell growth when the PAH compound is less bioavailable in the presence of hydrophobic surfaces (e.g. soil) (Wick et al. 2001; Wick et al. 2002a). Alternatively, PAH-degrading Mycobacterium and Sphingomonas bacteria could make use of other inorganic or organic nitrogen + - sources than the inorganic N-NH4 and N-NO3 that are available in the soil or the contaminant mixture. Some PAH- and oil-degrading bacteria have been reported to utilize aromatic organic sources of nitrogen and phosphorus such as for example heterocyclic aromatic azaarenes (Rosenberg et al. 1996; Willumsen et al. 2001b).

Imbalanced high CPAH/N/P-ratios following inorganic nutrient addition did not have a positive or negative effect on PAH-biodegradation by Mycobacterium and Sphingomonas strains. Similarly, an other study reported that in a soil that already had high natural N- and P- concentrations, the biodegradation of PAHs was not extra stimulated, nor depressed by the addition of an N+P-supplement with an excess of P creating a CPAH/N/P ratio of 120/12000/120000 [mg] (Johnson et al. 1990). In addition, a theoretical excess or lack of nitrogen added as ammonium (C/N = 100/70 or C/N = 100/3) or as nitrate (C/N=100/110 or C/N=100/2) did not influence the growth of LB126 on fluorene in chemostat experiments (van Herwijnen R. et al. unpublished). However, we discovered that addition of high concentrations of nitrogen or phosphorus supplements can totally block biodegradation by high increments of the salinity (I). Other researchers also observed that the N+P- supplement supplements inhibited phenanthrene degradation in three soils which became limited in nitrogen after P-excess amendment (N/P< 1) (Johnson et al. 1990). A significant difference between the effects caused by the addition of nitrogen salts, phosphorus salts or both has been reported in several studies (Swindoll et al. 1988; Manilal et al. 1991). As suggested before, nutrient additions may only be beneficial after the soils own available N- and P- reserves are depleted (Bossert et al. 1984). The knowledge regarding inorganic nutritional requirements for PAH-degrading Sphingomonas and Mycobacterium species obtained in this study could add to our limited understanding of their adapted physiology for PAHs degradation and could reveal ways to manipulate them successfully in order to enhance bioremediation of PAH-contaminated soils.

- 143 -

“One never notices what has been done; one can only see what remains to be done.”

- Marie Curie -

- 144 - General Discussion and Perspectives

CHAPTER 7

GENERAL DISCUSSION AND PERSPECTIVES

In bioremediation, soils contaminated with polycyclic aromatic hydrocarbons (PAHs) are often treated as a ‘black box’ without knowledge about the actual microbial catalysts involved in degradation. However, to optimize and control biodegradation processes this information is essential. Sphingomonas and Mycobacterium strains might be important PAH-degraders in PAH-contaminated soils as they have been often isolated in enrichments for PAH-degrading bacteria from such habitats. Moreover, PAH-degrading Sphingomonas and Mycobacterium strains seem to possess not only the appropriate enzymatic systems to degrade a wide variety of PAHs but also special mechanisms to get access to poorly bioavailable PAHs. Within the Mycobacterium genus, the capability to degrade PAHs is often found among strains related to the species M. gilvum and M. austroafricanum and especially M. frederiksbergense, a relatively recently described Mycobacterium species of which all members are PAH-degrading strains. Similarly, PAH-degrading Sphingomonas isolates are mainly members of the species of the former Sphingobium genus, i.e., S. yanoikuyae, S. xenophaga, S. chlorophenolicum, S. chungbukensis, S. herbicidivorans, and S. cloaca. Sphingomonas sp. EPA505, a model strain for the PAH-degrading Sphingomonas strains, forms phylogenetically possibly a new species together with some new PAH-degrading isolates grouped in a separated branch in the 16S rRNA gene based phylogenetic tree of the Sphingomonas genus.

However, current knowledge about PAH-degrading Mycobacterium and Sphingomonas strains is mainly based on cultivation–based isolated strains. Not much is know about the in situ occurrence, distribution and PAH-degradation activity of Sphingomonas and Mycobacterium strains and their specific nutrient requirements to become stimulated.

- 145 - Chapter 7

In the first part of our study, new genus-specific methods were developed to specifically detect PAH-degrading Mycobacterium and Sphingomonas species and to gain information about their diversity in soil. This was accomplished via specific PCR amplification of 16S rRNA genes followed by DGGE-analysis of the amplicons generating a Mycobacterium or Sphingomonas community fingerprint. The genus specific PCR-DGGE detection systems are indispensable tools to study the natural occurrence and diversity of Mycobacterium and Sphingomonas populations in soil. Moreover, the detection systems will allow to specifically monitor their dynamics during bioremediation and to further optimize bioremediation processes as the methods can be used to analyze whether or not certain control parameters positively or negatively affect Sphingomonas and Mycobacterium populations and if this is related to PAH-removal. Genus specific detection methods targeting a larger group of Mycobacterium or Sphingomonas species will exponentially amplify mainly the most abundant species, overshadowing the detection of other less abundant species. This was indeed shown by the detection of M. frederiksbergense and Sphingomonas sp. EPA505 related strains using two newly designed species-specific primers sets in soils in which these species were not detected using the genus specific primer sets. The strain specific PCR-based detection methods developed for these two species are thus valuable and can reveal additional information on the Mycobacterium and Sphingomonas population by detecting species which are otherwise missed.

For future application, it will be necessary to make the methods more quantitative either by ‘competitive PCR (cPCR)’ (Bjerrum et al. 2002) or ‘Quantitative Real Time PCR (QPCR)’ (Tell et al. 2003). Although real time PCR has advantages over competitive PCR, competitive PCR can still be of major interest and easy to accomplish. The DNA or cloned 16S rRNA genes of Mycobacterium or Sphingomonas strains generating easily distinguishable profiles on the DGGE-gel can be used as a competitor to quantify simultaneously different members of the Mycobacterium or Sphingomonas population by comparison of band intensities for the total fingerprint on a DGGE-gel (Felske et al. 1998). The methods allow further in situ investigation of the interactions, the function, the dispersal, the mobility and the colonization patterns of PAH-degrading Mycobacterium spp. and Sphingomonas spp. in the soil matrix. They will be essential to monitor the dynamics of these important PAH-degrading bacteria during bioremediation processes. In addition, our detection

- 146 - General Discussion and Perspectives methods can be combined with other primers targeting the catabolic messenger RNA to study the active population of Mycobacterium and Sphingomonas involved in PAH- degradation in soil. Only recently, several authors reported the cloning and sequencing of genes coding for the enzymes involved in PAH-degradation pathways in Sphingomonas species (Romine et al. 1999b; Pinyakong et al. 2003a) and Mycobacterium species (Khan et al. 2001; Krivobok et al. 2003). It is now clear from sequence comparison that the catabolic proteins involved in PAH-degradation in Mycobacterium and Sphingomonas species form separate families (Khan et al. 2001; Pinyakong et al. 2003a). This would allow designing new primer sets for specific PCR detection of the corresponding genes in the PAH-degrading isolates or environmental habitats.

In the past, many PAH-degrading Sphingomonas and Mycobacterium strains have been isolated from diverse PAH-contaminated environments. Our results, based on culture-independent detection techniques, confirmed for the first time clearly the simultaneous and ubiquitous occurrence of fast-growing Mycobacterium and Sphingomonas species in PAH-contaminated soils originating from different locations and characterized by different geological and chemical properties. The wide distribution and high abundance of Mycobacterium and Sphingomonas strains in general and more specific of strains related to M. frederiksbergense and Sphingomonas sp. EPA505 in PAH-contaminated soils revealed in this work, indicate these bacteria are important colonizers and possibly endemic pollutant degraders in such soils. As such, it seems that in most contaminated soils a natural potential for biodegradation is present and costly bioaugmentation procedures are unneeded. On the other hand, we do not have yet information if these detected strains are effectively involved in PAH-degradation. This can be investigated by the recent reported approach of ‘Stable Isotope Probing (SIP)’ (Manefield et al. 2002). In this approach, a 13C-labeled C-source, in our case 13C-PAHs, is added to the soil of interest. Actively PAH consuming bacteria will incorporate the 13C-label in their DNA which can be, after DNA-extraction, separated from 12C-DNA by means of gradient centrifugation. Using our PCR detection method, then the 13C-DNA can be examined to elucidate whether or not and which Mycobacterium and Sphingomonas species actively participate in the PAH-degradation process.

- 147 - Chapter 7

Our study revealed also a correlation between the occurrence and abundance of Mycobacterium and Sphingomonas species and the PAH-concentration profile of the soils (See summary table – Appendix). Particularly high concentrations of Sphingomonas were detected in soils containing high PAH-concentrations (1000 – 3000 mg kg-1) (mainly phenanthrene), such as the And and Barl soils used in this study. Mycobacterium cells were found in higher concentrations in soils with low organic carbon and PAH-concentrations than in soils containing relatively high PAH- concentrations. From other studies it is known that in soils such as Barl and And, a large fraction of the PAHs is relatively bioavailable. Our results may indicate different niches of Mycobacterium and Sphingomonas strains in PAH-contaminated soils. High concentrations of more bioavailable and more easily degradable PAHs such as phenanthrene may enrich mainly PAH-degrading Sphingomonas strains. Fast-growing Mycobacterium species may naturally be selected in PAH-polluted soil enriched in poorly bioavailable and highly recalcitrant higher molecular PAHs. This hypothesis fits with the recurrent isolation of mainly phenanthrene degrading Sphingomonas strains in aqueous enrichments, while Mycobacterium strains are mainly isolated using more hydrophobic PAHs such as pyrene or less bioavailable sorbed PAHs. Mycobacterium species and especially the M. frederiksbergense strains, maybe better adapted to harsh oligothrophic soil conditions as they have a low energy demand and make use of several PAH bioavailability-enhancing mechanisms such as high-affinity uptake systems and adhesion to the substrate (Wick et al. 2001; Wick et al. 2002a; Miyata et al. 2004). In future experiments, this interesting observation can be further analyzed by for example monitoring Sphingomonas and Mycobacterium dynamics in time in soils containing low initial concentrations of bioavailable PAHs (e.g. soils B101, TM and E6068) that are supplemented with ‘easily available’ phenanthrene. In addition, it would be interesting to monitor community dynamics during removal of phenanthrene from soils such as And or Barl.

Further analysis of the biodiversity of Mycobacterium and Sphingomonas populations in PAH-contaminated soils, revealed a clear correlation between the Sphingomonas diversity and the PAH-concentration of the soils. A diverse group of Sphingomonas strains belonging to different species clusters in the genus was present in low and moderated contaminated soils at relatively equal cell concentrations. Soils containing high PAH-concentrations (mainly phenanthrene) showed less heterogeneous DGGE

- 148 - General Discussion and Perspectives community fingerprints as the Sphingomonas populations in these soils were probably dominated by high numbers of a few Sphingomonas strains which possibly masked the detection of other species that are present in lower concentrations. Sequence analysis of Sphingomonas communities from contaminated soils revealed the presences of ‘new’ 16S rRNA gene sequences grouped in possible new Sphingomonas species. Most soil extracted Sphingomonas sequences were only limited related to identified species and cultivated PAH-degrading isolates and probably represent truly ‘non-culturable’ Sphingomonas strains present in soil. Using a strains specific primer set, Sphingomonas sp. EPA505 related cells were detected in relative high concentrations in soils with both low and high concentrations of PAHs. Although, EPA505 related cells had never been found among the clone sequences retrieved from any of these soils with this Sphingomonas specific primer set or were never recovered from the same soils using a culture-based selective plating method (Vanbroekhoven et al. unpublished).

There was no clear correlation between Mycobacterium biodiversity (assessed by the number of bands in the Mycobacterium DGGE fingerprints) and the PAH- concentration of the soils. Sequences related to known PAH-degrading species such as M. frederiksbergense and M. austroafricanum but especially to M. tusciae were repeatedly detected in all PAH-contaminated soils, originating from different countries and different industrial sites, but not in the non-contaminated soil. Although, M. tusciae is not known to display PAH-degradation properties or has never been isolated as a PAH-degrader, the M. tusciae sequences isolated in this study grouped with other unidentified Mycobacterium sequences cloned from DNA from petroleum contaminated soils (Cheung et al. 2001). These results may indicate a so far unidentified but an important role for M. tusciae and/or related species in PAH- degradation processes in soil. Using the newly developed genus specific primer set, M. frederiksbergense related sequences were detected only in 1 soil, while using a newly developed M. frederiksbergense specific primer set M. frederiksbergense strains were detected additionally in 2 other soils with relatively low concentrations of PAH concentrations (< 100 mg kg-1) or high concentrations of oil (4600 mg kg-1). The role of individual species such as M. frederiksbergense or M. tusciae in PAH- contaminated soil can be further explored by ribosomal RNA analysis with RT-PCR techniques selecting for the active metabolizing population of a Mycobacterium or

- 149 - Chapter 7

Sphingomonas population. Moreover, in future experiments, also the contribution of different species in the PAH-degradation process can be assessed by ‘Stable Isotope Probing (SIP)’ (Manefield et al. 2002), as described above.

As carbon/nitrogen/phosphate nutrient conditions in untreated contaminated soils are mostly far from optimal, ‘biostimulation’ or the addition of nutrient supplements is a commonly used practice to stimulate bioremediation processes. However, not much is known about the nutrient requirements of PAH-degrading Mycobacterium and Sphingomonas strains in PAH-contaminated soil. Therefore, in a second part of our study, inoculated microcosm tests were set up to determine of the impact of bioavailable mineral nutrient concentration on the biodegradation activity of some model PAH-degrading Sphingomonas spp. and Mycobacterium spp.. It was demonstrated that PAH-degradation by Sphingomonas and Mycobacterium strains in soil slurry was found maximal under carbon/nitrogen/phosphate concentrations of circa 120/14/3 [expressed in mg] or 100/10/1 [expressed in mole], i.e., a ratio theoretical predicted as optimal for cell growth in general. Under these optimal nutrient conditions, metabolisation of PAHs sustained new biomass production by Sphingomonas and Mycobacterium cells. Mineralization curves were sigmoid in nature, which indicates that a period of growth was needed before significant mineralization of the added PAH occurred. Significant PAH removal was detectable when cell concentrations reached 106 cells per gram of soil.

In addition, PAH-biodegradation by inoculated Sphingomonas and Mycobacterium spp. was not constrained by imbalanced CPAH/N/P-ratios with low nitrogen and/or phosphate concentrations naturally occurring in most contaminated soils. It has been reported before that Mycobacterium cells can shift to ‘maintenance metabolism’ during PAH-degradation (Wick et al. 2001; Wick et al. 2002a; Miyata et al. 2004), indicating that they would need less nitrogen and phosphorus because more PAH carbon is used for energy production instead of biomass production than normally for other carbon substrates. Alternatively, PAH-degrading Mycobacterium and Sphingomonas bacteria could make use of other inorganic or organic nitrogen sources + - than the inorganic N-NH4 and N-NO3 that are available in the soil.

- 150 - General Discussion and Perspectives

In general it is believed that nutrient amendments will enhance survival and biodegradation activity of the soil bacteria, increasing biodegradation rates and reducing bioremediation costs. However, a significant difference between the effects caused by the addition of nitrogen salts, phosphorus salts or both has been reported in many bioremediation studies. In our study, imbalanced high CPAH/N/P-ratios following inorganic nutrient addition did not have a positive or negative effect on PAH-biodegradation by Mycobacterium and Sphingomonas strains. However, we discovered that addition of high concentrations of nitrogen or phosphorus supplements can totally block biodegradation by high increments of the salinity (I). Nutrient additions may only be beneficial after the soils own available N- and P- reserves are depleted. Thus adding blindly large amounts of nutrients to support or stimulate biodegradation may affect biodegradation negatively.

Our results concerning occurrence and nutrient demands of PAH-degrading Mycobacterium species in PAH-contaminated soils support the hypothesis that PAH- degrading Mycobacterium strains are ‘K-strategists’, i.e., relatively slow growing bacteria adapted to a resource restricted and crowded environment, like it has been suggested for their close relatives Rhodococcus and Pseudonocardia (Juteau et al. 1999). Also PAH-degrading Sphingomonas species show characteristics of ‘K- strategists’ (relatively slow growing). On the other hand, PAH-degrading bacteria such as Pseudomonas species for example, could be considered as ‘r-strategists’, i.e., relatively fast growing bacteria adapted to a resource-abundant and uncrowded environment. The main advantage of r-strategists is their high growth rates while K- strategists are characterized by other competitive abilities such a high affinity for the substrate (probably most important) or resistance to predation (shown for Rhodococcus and Pseudonocardia) (Juteau et al. 1999). This r/K selection theory (Andrews et al. 1986) could explain why Sphingomonas and especially Mycobacterium species are not outnumbered by faster growing PAH-degrading bacteria such as Pseudomonas species for example. PAH-contaminated soil and its Sphingomonas-Mycobacterium community fit the description of a K-environment very well. Mycobacterium and Sphingomonas species are certainly a very important group of PAH-degrading bacteria well adapted to the harsh environmental conditions and with clearly great capacities for biorestauration of contaminated sites.

- 151 - Chapter 7

The results and conclusions of the 4 year research presented in this thesis will be indispensable for future studies concerning biodegradation of PAHs by Sphingomonas and Mycobacterium in the environment. The knowledge regarding inorganic nutritional requirements for PAH-degrading Sphingomonas and Mycobacterium species obtained in this study expands our understanding of their adapted physiology for PAHs degradation and could reveal ways to manipulate them successfully in order to enhance biodegradation in PAH-contaminated soils. For further bioremediation optimization, the detection techniques developed in this study are currently used by different research groups to study the dynamics of Mycobacterium and Sphingomonas communities in response to various bioremediation technologies and set-ups. The influence of control parameters such as oxygen and nutrient supply on the behavior and PAH-degrading activity of the Mycobacterium/Sphingomonas spp. is evaluated. Finally, this research may enable us to improve bioremediation technologies so that we can stimulate as efficiently as possible the key biodegrading organisms and particularly fulfill their requirements. It will aid the development of a bioremediation technology which is better understood, better predictable, better controllable and more efficient on a large-scale.

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- 176 - Appendix.

APPENDIX

Summary table of all tested soils and the different PCR-DGGE results.

- 177 - Appendix.

MYCO MYCO† MYCOF†† SPHINGO° EPA°° Soil TOC PAH conc. Oil conc. DNA conc.* Nested‡ Soil Origin pH (estimated (estimated (estimated (estimated type (%) (mg kg-1) (mg kg-1) (µg g-1) (estimated cells g-1) cells g-1) cells g-1) cells g-1) cells g-1) S587 Corn field (Belgium) sand 5.5 2.15 0.289 < 50 31.00 + ND ND ND ND S588 Horse pasture (Belgium) sand 6.0 2.46 0.391 < 50 18.00 + ND ND ND ND S585 Pine tree forest (Belgium) sand 5.8 3.19 0.673 < 50 31.75 + ND ND ND ND S589 Ditch in agricultural area (Belgium) sand 5.8 4.24 0.721 < 50 49.50 + ND ND ND ND S592 Vegetable garden (Belgium) sand 7.0 3.16 1.011 < 50 38.25 + ND ND ND ND S584 Compost heap (Belgium) sand 7.3 7.04 1.063 < 50 27.75 + ND ND ND ND S591 Non-paved land road (Belgium) sand 9.0 0.76 3.357 < 50 6.25 NP ND ND ND ND TB3 Coal gasification plant (Belgium) sand 8.23 1.52 14 < 50 2.65 + + ND ND ND K3840 Gasoline station site (Denmark) Sand 8.20 0.50 20 98 2.75 + (109) + + (109) + (106) + (106) B101 Coal gasification plant (Belgium) Sand 7.00 2.63 107 70 27.25 + (108) + + (108) + (105) + (104) E6068 Gasoline station site (Denmark) Sand 7.96 9.94 258 300 5.40 + (107) + NP (<106) ND ND TM Coal gasification plant (Belgium) Sand 8.00 3.85 506 4600 4.75 + (108) + + (106) + (106) + (105) Barl Coal gasification plant (Germany) Gravel 8.90 4.63 1029 109 6.15 NP (<106) NP (<102) NP (<106) + (106) NP (<104) AndE Railway station site (Spain) Clay 8.10 2.35 3022 2700 3.40 NP (<106) + NP (<106) + (106) + (106)

* DNA recovery per g soil, mean value of 2 parallel extractions of 1 gr of soil. † result of direct PCR on soil DNA extract with Mycobacterium specific primers MYCO66f and GC40-MYCO600r. ‡ result of nested PCR on soil DNA extract with eubacterial primers 27f and 1492r followed by Mycobacterium specific primers MYCO66f and GC40-MYCO600r. †† result of direct PCR on soil DNA extract using M. frederiksbergense specific primers MYCOFf and MYCOFr. ° result of direct PCR on soil DNA extract with Sphingomonas specific primers Sphingo108f & GC40-Sphingo420r. °° result of direct PCR on soil DNA extract with Sphingomonas sp. EPA505 species specific primers EPAf and EPAr. Symbols: + = detectable PCR product, NP = no detectable PCR product in agarose electrophoresis of PCR products, ND = not determined. Estimated cells g-1 = roughly estimated cell content based on a ‘dilution to extinction’ PCR approach using only specific primers are indicated between brackets.

.

- 178 - Notation Index.

NOTATION INDEX

bp base pairs (DNA size indicator) C-source compound is used as sole source of carbon DGGE denaturing gradient gel electrophoresis DNA deoxyribonucleic acid EPS extra polysaccharides HGT horizontal gene transfer HMW-PAH high molecular weight PAH I ionic strength ISP iron suphur protein LMW-PAH low molecular weight PAH MGE mobile genetic elements NAPL non-aqueous phase liquid N-source compound is used as sole source of nitrogen OC organic carbon PAH polycyclic aromatic hydrocarbon PCR polymerase chain reaction P-source compound is used as sole source of phosphorus RNA ribonucleic acid RT-PCR reverse transcription polymerase chain reaction SOM soil organic matter

- 179 -

“Prediction is very difficult, especially about the future.”

- Niels Bohr -

- 180 - Curriculum Vitae

CURRICULUM VITAE

Ir. Natalie M.E.J. Leys

Nationality Belgian Date & place Birth °23/3/1975, Lier, Belgium Sex Female Identity card No. 067-0017005-78 Register No. 750323-118-12

Co-ordinates

Private

Postal address Fazantenlaan 20, 2290 Vorselaar, Belgium Phone +32-(14)-514224 or +32-(473)-505286

Business

Position Scientific Collaborator Belgian Nuclear Research Centre SCK•CEN Division Radioactive Waste & Clean Up Laboratory of Microbiology Postal address Boeretang 200, 2400 Mol, Belgium Phone +32-(14)-332726 Mail [email protected]

Professional Employment

Oct. 2002 - today Scientific project manager Laboratory of Microbiology, Section Radioactive Waste and Clean-up, Belgian Nuclear Research Centre (SCK●CEN), Mol, Belgium

Oct. 1998 – Oct. 2002 PhD researcher University of Ghent (UG), Ghent, Belgium & Flemish Institute of Technological Research (Vito), Mol, Belgium

Curriculum Vitae

Education

Scientific Academic Training

1998 - 2004 Ph.D. in Applied Biological Sciences In preparation University of Gent (UG), Ghent, Belgium PhDThesis: “PAH-biodegradation by Sphingomonas and Mycobacterium : their occurence, diversity and nutrient demands in PAH contaminated soils” Performed at the Univerity of Gent (UG), Gent, Belgium and Flemisch Institute for Technological Research (Vito), Mol, Belgium. Scientific promotors UG: Prof. W. Verstraeten and Prof. E. Top Scientific promotors Vito: Dr. D. Springael and Dr. L. Bastiaens

1995 - 1998 Educational degree in Biology (Aggregation) Cum laude Katholic University of Leuven (KUL), Leuven, Belgium

1995 - 1998 Master of Bio-engineer in Chemistry, Option Industrial Microbiology, Cum laude Katholic University of Leuven (KUL), Leuven, Belgium Master's Thesis: "Melkzuurfermentaties in gefermenteerd maïsmeel en sorghum

bier”, Stellenbosch University, South-Africa, Scientific Promotor KUL: Prof. H. Verachtert

1993 - 1995 Bachelor of Bio-engineer in Chemistry, Cum laude Katholic University of Leuven (KUL), Leuven, Belgium

1987 - 1993 High school, section Latin-Sciences Cum laude Kardinaal Van Roey Institute, Vorselaar, Belgium

Participation in Research projects

2002 - 2004 Principal Scientific Researcher of the ‘MESSAGE’ Space Research Project ESA project MESSAGE: Microbial Experiment in Space Station About Gene Expression MESSAGE 1 experiment, Belgian Taxi Flight, 30 October – 10 November 2002 MESSAGE 2 experiment, Spanish Soyuz Mission, 18 – 28 October 2003 Through participation of the Belgian Nuclear Research Centre (SCK), Mol, Belgium 2000 – 2002 Member of the research team of the ‘Biostimul’ European Project, Through participation of the Flemish Institute of Technological Research (Vito), Mol, Belgium 1998 - 2000 Member of the research team of the ‘Biovab’ European Project, Through participation of the Flemish Institute of Technological Research (Vito), Mol, Belgium

Curriculum Vitae

Publications

Publications in International journals with referee

Springael, D., Vanbroekhoven, K., Leys, N., Bastiaens, L., Ryngaert, A, Vancanneyt, M, Swings, J., Wattiau, P. and Diels. L., Isolation of PAH degrading bacteria using habitat directed methods. FEMS Microbiol. Ecol., In preparation.

Leys, N., Bastiaens, L., Springael, D. Bacterial Biodegradation of PAHs in contaminated habitats: A literature Review, Curr. Adv. Appl. Microbiol.Biotechnol. Submittted.

Leys, N., Bastiaens, L., Springael, D. Microbial bioremediation of PAH-contaminated soil: A literature Review, Curr. Adv. Appl. Microbiol.Biotechnol. Submittted.

Leys N., Bastiaens L., W. Verstraete, D. Springael, Influence of the Carbon/Nitrogen/Phosphate-ratio in soil on PAH-degradation by Mycobacterium and Sphingomonas strains, Appl. Microbiol. Biotechnol., Submitted.

Leys N., Ryngaert A., Bastiaens L., van Canneyt Mark, Swings Jean,E. Top, W. Verstraete, D. Springael, Mycobacterium frederiksbergens, specialised in degradation of polycyclic aromatic hydrocarbons, is ubiquiteous in PAH-contaminated soil, Environ. Microbiol., Submitted.

Leys N., Ryngaert A., Bastiaens L., E. Top, W. Verstraete, D. Springael, Occurrence of Sphingomonas sp. EPA505 related strains in soils contaminated with polycyclic aromatic hydrocarbons (PAHs)., Microbiol. Ecol., Submitted.

Leys N., Ryngaert A., Bastiaens L., P. Wattiau, E. Top, W. Verstraete, D. Springael, Occurrence and phylogenetic diversity of fast-growing Mycobacterium species in PAH-contaminated soils. FEMs Microbiol. Ecol., Submitted.

Leys N., Ryngaert A., Bastiaens L., E. Top, W. Verstraete, D. Springael, (2004) Occurrence and phylogenetic diversity of Sphingomonas in PAH-contaminated soils., Appl. Environ. Microbiol. 70(4): 1944-1955.

Publications in books and proceedings without referee

Leys N., Bastiaens L., Wattiau P., Vanbroekhoven K., Gemoets J., Top E., Springael D., 1999, Detection of consortia of PAH-degrading bacteria in soil by PCR using strain- and genus specific primers combined with DGGE and SCCP, In: Proceedings of the 13th Forum for Applied Biotechnology, 22-23 September 1999. Mededelingen Faculteit Landbouwkundige en Toegepaste Biologische Wetenschapppen, 64 (5a), 155-158.

Vanbroekhoven K., Bastiaens L., Leys N., De Mot R., Springael D., 1999, Environmental monitoring of bioavailability promoting strains in a LNAPL contaminated soil, In: Proceedings of the 13th Forum for Applied Biotechnology, 22-23 September, 1999. Mededelingen Faculteit Landbouwkundige en Toegepaste Biologische Wetenschapppen, 64 (5a), 215-218.

Curriculum Vitae

Participation to International Congresses and Symposia

International congress with oral presentation

Springael D., Leys N., Bastiaens L., Wattiau P., Top E., 2002, Monitoring of PAH-degrading Mycobacterium spp. strains in soil samples. International Symposium on Subsurface Microbiology, Copenhagen, Denmark.

Leys N., 2000, PCR combined with Denatured Gradient Gel Electrophoresis (DGGE) or Single Strand Conformation Polymorphism (SSCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples, Meeting of the Section Microbial Ecology, NIOO- Centre for Terrestrial Ecology, Heteren, The Netherlands.

International congress with poster presentation

Leys N., R. Wattiez, S. Baatout, P. De Boever, M. Mergeay, 2003, Gene expression in Ralstonia metallidurans CH34 in space., European Conference on Prokaryotic Genomes,Göttingen, Germany, October 2003

Leys N., Bastiaens L., Wattiau P., Top E., Springael D., 2001, Monitoring of PAH-degrading Sphingomonas spp. and Mycobacterium spp. in soil samples. (P.23.019) 9the International Symposium on Microbial Ecology (ISME9), Amsterdam, The Netherlands.

Leys N., Bastiaens L., Weurtz S., Top E., Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) to specifically detect and monitor PAH-degrading Sphingomonas spp. in environmental samples. Symposium on emerging and recurrent infectious diseases, Louvain-La-Neuve, Belgium.

Leys N., Van Broekhoven K., Bastiaens L., Wattiau P., Top E., Tebbe C, Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) or Single Strand Conformation Polymorphism (SSCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples. Symposium on emerging and recurrent infectious diseases, Louvain-La- Neuve, Belgium.

Leys N., Van Broekhoven K., Bastiaens L., Wattiau P., Top E., Tebbe C, Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) or Single Strand Conformation Polymorphism (SSCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples. Theme Day: New Trends in soil Microbial Ecology, ULB, Brussels, Belgium.

Leys N., Bastiaens L., Weurtz S., Top E., Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) to specifically detect and monitor PAH-degrading Sphingomonas spp. in environmental samples. Theme Day: New Trends in soil Microbial Ecology, ULB, Brussels, Belgium.

Springael D., Van Broekhoven K., Bastiaens L., Ryengaert A., Leys N., Wattiau P., Diels L., 2000, PCR combined with Denaturing Gradient Gel Comparative study of different isolation protocols providing Polycyclic Aromatic Hydrocarbons (PAHs) in various available states in order to isolate novel PAH-degraders. Secterial Meeting for EC Project Contractors Bioremediation: Biotechnology for Environmental Applications, Roskilde, Denmark.

Leys N., Van Broekhoven K., Bastiaens L., Wattiau P., Top E., Tebbe C, Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) or Single Strand Conformation Polymorphism (SSCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples. Secterial Meeting for EC Project Contractors Bioremediation: Biotechnology for Environmental Applications, Roskilde, Denmark.

Curriculum Vitae

Leys N., Bastiaens L., Weurtz S., Top E., Springael D., 2000, PCR combined with Denaturing Gradient Gel Electrophoresis (DGGE) to specifically detect and monitor PAH-degrading Sphingomonas spp. in environmental samples. Secterial Meeting for EC Project Contractors Bioremediation: Biotechnology for Environmental Applications, Roskilde, Denmark.

Leys N., Vanbroekhoven K., Bastiaens L., Wattiau P., Top E., Tebbe C., Springael D.,, 2000, PCR combined with denaturing gradient gel electrophoresis (DGGE) or single strand conformation polymorphism (SCCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples, 4the International Symposium on Environmental Biotechnolgy (ISEB4), Noordwijkerhout, The Netherlands.

Leys N., Vanbroekhoven K., Bastiaens L., Wattiau P., Top E., Tebbe C., Springael D., 2000, PCR combined with denaturing gradient gel electrophoresis (DGGE) or single strand conformation polymorphism (SCCP) to specifically detect and monitor PAH-degrading Mycobacterium spp. in environmental samples, International Workshop on Methods to monitor microbial inoculants to improve their success, Wageningen, The Netherlands.

Vanboekhoven K., Bastiaens L., Leys N., De Mot R., Springael D., 1999, Evironmental monitoring of bio-availability promoting strains in LNAPL contaminated soil. Symposium on Microbial identification: an integrative approach, Gasthuisberg, Leuven, Belgium.

Springael D., Vanbroekhoven K., Bastiaens L., Ryngaert A., Leys N., Wattiau P., Diels L., 1999, Comparative study of different isolation protocols which provide Polycylcic Aromatic Hydrocarbons (PAHs) in various available states inorder to isolate novel PAH-degrading bacteria, Symposium on Microbial identification: an integrative approach, Gasthuisberg, Leuven, Belgium.

Leys N., Bastiaens L., Wattiau P., Vanbroekhoven K., Gemoets J., Top E., Springael D., 1999, Detection of consortia of PAH-degrading bacteria in soil by PCR using strain- and genus specific primers combined with DGGE, Symposium on Microbial identification: an integrative approach, Gasthuisberg, Leuven, Belgium.

Vanboekhoven K., Bastiaens L., Leys N., De Mot R., Springael D., 1999, Evironmental monitoring of bio-availability promoting strains in LNAPL contaminated soil. 13 the Forum for Applied Biotechnology (FAB99), Gent, Belgium.

Leys N., Bastiaens L., Wattiau P., Vanbroekhoven K., Gemoets J., Top E., Springael D., 1999, Detection of consortia of PAH-degrading bacteria in soil by PCR using strain- and genus specific primers combined with DGGE and SCCP, 13 the Forum for Applied Biotechnology (FAB99), Gent, Belgium.

Leys N., Bastiaens L., Wattiau P., Vanbroekhoven K., Gemoets J., Top E., Springael D., 1999, Detection of consortia of PAH-degrading bacteria in soil by PCR using strain- and genus specific primers combined with DGGE and SCCP, 13 the Forum for Applied Biotechnology (FAB99), Ghent, Belgium.