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Two novel transporters essential for the reassimilation of cholic acid metabolites excreted by Rhodococcus.. Swain, Kendra Lynn 2011

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TWO NOVEL TRANSPORTERS ESSENTIAL FOR REASSIMILATION OF CHOLIC ACID METABOLITES EXCRETED BY Rhodococcus jostii RHA1 by Kendra Lynn Swain  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in  THE FACULTY OF GRADUATE STUDIES (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  August 2011  © Kendra Lynn Swain, 2011  Abstract  In this study, I investigated two novel transporters associated with cholic acid catabolism in Rhodococcus jostii RHA1. Reverse-transcriptase quantitative-PCR indicated that an ABC transporter was upregulated 16.7-fold and an MFS transporter was upregulated 174-fold during the exponential phase of growth on cholic acid compared to pyruvate. With gene knockout analysis, I discovered that these transporters are required for the reassimilation of distinct cholic acid metabolites. The ABC transporter, encoded by the camABCD genes, was essential for uptake of 12-hydroxy-9-oxo-1,2,3,4,10,19,23,24octanorcholan-5,22-dioic acid and 12-hydroxy-9-oxo-1,2,3,4,10,19,23,24-octanorchol-6-en5,22-dioic acid. The MFS transporter, encoded by the camM gene, was essential for uptake of 3,7,12-trihydroxy-9-oxo-9,10-seco-23,24-bisnorchola-1,3,5(10)-trien-22-oic acid. The uptake of these metabolites is necessary for maximal growth on cholic acid: the camB mutant, lacking the permease component of the ABC transporter, and the camM mutant, lacking the MFS transporter, only achieved 74% and 77%, respectively, of the final wild type protein yield. These metabolites differ from previously reported cholic acid metabolites from Proteobacteria in that they retain an isopropionyl side chain at the C17 position. This study is the first to demonstrate the function of putative cholic acid genes through targeted mutagenesis, as well as the first to provide evidence for the requirement for transporters involved in cholic acid metabolite uptake. This work highlights the importance and complexity of transport processes associated with bacterial catabolism and may contribute to industrial applications involving bacterial steroid transformation.  ii  Table of Contents  Abstract .................................................................................................................................... ii Table of Contents ................................................................................................................... iii List of Tables ......................................................................................................................... vii List of Figures ....................................................................................................................... viii Acknowledgements ................................................................................................................ xi Chapter 1: Introduction ........................................................................................................ 1 1.1  Steroids ..................................................................................................................... 1  1.1.1  Steroid Function .................................................................................................... 1  1.1.2  Steroid Therapeutics ............................................................................................. 2  1.1.3  Steroids in the Environment.................................................................................. 2  1.2  Microbial Steroid Degradation ................................................................................. 4  1.2.1  Harnessing Microbial Steroid Catabolic Pathways for Industrial Purposes ......... 4  1.2.2  Steroid Bioremediation using Microorganisms .................................................... 6  1.3  The Seco-steroid Degradation Pathway .................................................................... 7  1.3.1  Microbial Testosterone Degradation..................................................................... 8  1.3.1.1  17-Dehydrogenation ................................................................................. 10  1.3.1.2  Transformation of Androstenedione ........................................................... 10  1.3.1.3  Transformation of DOHNAA ..................................................................... 11  1.3.1.4  Steroid Pathway Gene Clusters in C. testosteroni ...................................... 12  1.3.2  Microbial Cholesterol Degradation..................................................................... 12  1.3.2.1  Modeling M. tuberculosis Cholesterol Degradation in R. jostii RHA1 ...... 12  iii  1.3.2.2  Cholesterol Uptake...................................................................................... 13  1.3.2.3  Cholesterol Side-Chain Degradation and A-ring Oxidation ....................... 13  1.3.2.4  Predicting Cholesterol Pathway Gene Function ......................................... 16  1.3.2.5  Rings A and B Degradation and DOHNAA Generation ............................ 16  1.3.3  Microbial Bile Salt Degradation ......................................................................... 17  1.3.3.1  Bile Salt Degradation by Free-Living Bacteria .......................................... 18  1.3.3.2  Relevance of Microbial Bile Salt Transformation ...................................... 19  1.3.3.3  The Unifying Scheme of Microbial Bile Salt Degradation ........................ 20  1.3.3.4  Initiation of the Bile Salt Pathway .............................................................. 20  1.3.3.5  -Oxidation of Bile Salts in Pseudomonas sp. Strain Chol1 ........................ 21  1.3.3.6  Degradation of Rings A and B .................................................................... 22  1.4  RHA1: An Excellent System to Study Cholic Acid Catabolism ............................ 24  1.4.1  The Unique Properties of Actinobacteria ........................................................... 24  1.4.2  The Resilience of Rhodococci ............................................................................ 24  1.4.3  Bile Salt Pathway Genes in RHA1 ..................................................................... 25  1.4.4  The Aim of this Study ......................................................................................... 27  Chapter 2: Materials and Methods .................................................................................... 29 2.1  Reagents and Chemicals ......................................................................................... 29  2.2  Cell Growth Assays ................................................................................................ 29  2.3  DNA Manipulation and Construct Preparation....................................................... 31  2.4  Cloning of Knockout Mutants ................................................................................ 35  2.5  Cloning of Complementation Strains...................................................................... 36  2.6  RNA Extraction ...................................................................................................... 38  iv  2.7  cDNA Synthesis ...................................................................................................... 39  2.8  RT-QPCR Analysis ................................................................................................. 39  2.9  Chemical Analysis .................................................................................................. 40  Chapter 3: Results................................................................................................................ 42 3.1 3.1.1  RHA1 Cholic Acid Growth Phenotype................................................................... 42 Extracellular Cholic Acid Metabolite Accumulation by RHA1 ......................... 44  3.2  Expression of the ATP-Binding Cassette and MFS Transporters .......................... 45  3.3  Inactivation of the ABC Transporter ...................................................................... 48  3.3.1  The Cholic Acid Growth Phenotype of camB .................................................. 49  3.3.2  Extracellular Cholic Acid Metabolite Accumulation by camB ........................ 50  3.3.3  RHA1 and camB Growth on HOONCA Metabolites ...................................... 54  3.4  Inactivation of the MFS Transporter....................................................................... 55  3.4.1  The Cholic Acid Growth Phenotype of camM ................................................. 55  3.4.2  Extracellular Cholic Acid Metabolite Accumulation by camM ....................... 57  3.4.3  RHA1 and camM Growth on THOSBNC ........................................................ 59  3.5  Cholic Acid Conjugate Growth Phenotype of camM........................................... 60  3.6  Complementation .................................................................................................... 61  Chapter 4: Discussion .......................................................................................................... 63 4.1  Cholic Acid Uptake and the Potential for Passive Diffusion .................................. 63  4.2  A Unifying Scheme for Steroid Degradation.......................................................... 64  4.3  Support for the Toxic Metabolite Excretion Hypothesis ........................................ 65  4.4  Two Novel Transporters ......................................................................................... 67  4.5  Habitat of Cholic Acid Degraders........................................................................... 67  v  4.6  Implications............................................................................................................. 68  References .............................................................................................................................. 70  vi  List of Tables  Table 2.1 Stock salt solution recipe......................................................................................30 Table 2.2 Concentrated Goodies recipe.................................................................................30 Table 2.3 100X Mg/Ca/B1/Goodies mix recipe....................................................................31 Table 2.4 M9 basal salts recipe for 1L.................................................................................31 Table 2.5 Fosmids and plasmids used in this study................................................................32 Table 2.6 Oligonucleotides used in this study.........................................................................34  vii  List of Figures Figure 1.1 Stucture of the steroid nucleus: cyclopentanoperhydrophenantrene (sterane)........1 Figure 1.2 -Sitosterol and cholesterol; the two most common starter steroids for whole cell bioconversion.............................................................................................................................5 Figure 1.3 Proposed testosterone degradation pathway of C. testosteroni TA441...................9 Figure 1.4 Proposed cholesterol side-chain degradation pathway..........................................14 Figure 1.5 Deduced cholesterol nucleus degradation pathway...............................................15 Figure 1.6 Structure of cholic acid..........................................................................................17 Figure 1.7 Proposed -oxidation of the cholic acid side chain...............................................21 Figure 1.8 Structure of DHOPDC...........................................................................................22 Figure 1.9 Structure of DHADD.............................................................................................22 Figure 1.10 Structure of THSATD.........................................................................................23 Figure 1.11 ATP-binding cassette transporter........................................................................26 Figure 1.12 Multi-drug transporter of the MFS superfamily..................................................26 Figure 3.1 Growth of RHA1 on 2.0 mM cholic acid as the sole carbon and energy source..42 Figure 3.2 Final protein assay of all strains grown on 2.0 mM cholic acid............................43 Figure 3.3 Cholic acid metabolite HOONCA.........................................................................44 Figure 3.4 Cholic acid metabolite 6-HOONCA....................................................................44 Figure 3.5 Cholic acid metabolite THOSBNC.......................................................................45 Figure 3.6 Relative upregulation of camB during growth on cholic acid based on RT-QPCR.................................................................................................................................46 Figure 3.7 The ABC-transporter operon genes.......................................................................47  viii  Figure 3.8 Relative upregulation of camM during growth on cholic acid based on RTQPCR.......................................................................................................................................47 Figure 3.9 Colony PCR electrophoresis gel screening for camB transconjugation.............48 Figure 3.10 camB cholic acid phenotype compared to wild type........................................49 Figure 3.11 Accumulation of HOONCA metabolites in wild type RHA1 versus camB, based on GCMS normalized peak abundance.........................................................................51 Figure 3.12 Chromatogram of camB stationary phase supernatant showing peaks of accumulated metabolites..........................................................................................................52 Figure 3.13 Mass spectrum of HOONCA...............................................................................52 Figure 3.14 Mass spectrum of 6-HOONCA..........................................................................53 Figure 3.15 Color-comparison of supernatants.......................................................................53 Figure 3.16 Growth of RHA1 and camB on HOONCA metabolites as a carbon and energy source.......................................................................................................................................54 Figure 3.17 Colony PCR electrophoresis gel screening for camM transconjugation..........55 Figure 3.18 Growth of camM on 2.0 mM cholic acid, compared to wild type measured via OD600.................................................................................................................56 Figure 3.19 Cholic acid growth curve measured as protein....................................................57 Figure 3.20 THOSBNC accumulation in RHA1 and camM based on normalized GCMS peak abundance........................................................................................................................58 Figure 3.21 Chromatogram of camM stationary phase culture showing peaks of accumulated metabolites..........................................................................................................58 Figure 3.22 Mass Spectrum of THOSBNC............................................................................59  ix  Figure 3.23 Growth of RHA1 and camM on the THOSBNC metabolite as a carbon and energy source...........................................................................................................................60 Figure 3.24 Structures of conjugated bile salts.......................................................................61  x  Acknowledgements  I would like to thank my supervisor, Dr. William W. Mohn, for his wisdom, insight, and genuine kindness as he guided me through my journey as a graduate student. Dr. Mohn encouraged me to develop as an independent thinker and a capable scientist.  I thank my committee members, Dr. Erin Gaynor and Dr. Lindsay Eltis, for their intelligent perspective and thoughtful suggestions, as well as their positive affirmations when I was on the right track.  I appreciate my lab members for creating a hard-working, amiable environment that made it a joy to work in our lab. I’d particularly like to thank Justin LeBlanc for training me in RTQPCR, and for holding my hand through the trials of the SacB protocol. I appreciate Dr. Somalinga Vijayakumar for being there to bounce ideas off of when I needed a second opinion. I especially thank Gordon Stewart for being my “go to” guy for random bits of information, and for all the help he provided me with the GCMS.  I feel very grateful for the help of Jie Liu with all the cloning involved in my project, especially for teaching me her “fool proof” method for designing primers. I also appreciate Dr. Israel Casabon for helping me name the cholic acid metabolites I isolated.  Finally, I’d like to thank Alex Schwartz for believing in me and emotionally supporting me through the most stressful periods of being a graduate student.  xi  Chapter 1: Introduction  1.1  Steroids  Steroids are a class of organic compounds composed of a nucleus of 17 carbon atoms, arranged in 3 cyclohexane rings and a cyclopentane ring fused together, usually referred to as rings A through D (Figure 1.1). Steroids vary in structure by functional groups bound to different sites on this steroid nucleus; there are hundreds of variations found in animals, plants and fungi [1].  Figure 1.1 Structure of the steroid nucleus: cyclopentanoperhydrophenantrene (sterane).  1.1.1  Steroid Function  Different kinds of steroids perform different functions. Within animals, there are the hormone steroids, which are responsible for sexual differentiation, reproduction, regulation of metabolism, immune function, and muscle and bone synthesis to name a few [2]. As well, cholesterol is a major mammalian steroid responsible for modulating fluidity of cell membranes. Ergosterol in fungi and phytosterols in plants essentially perform the same 1  membrane function as cholesterol in mammals [3]. There are only a few reports of steroid synthesis in bacteria; including certain methanotrophic bacteria, myxobacteria, and marine bacteria [4].  1.1.2  Steroid Therapeutics  Steroids are also major pharmaceutical compounds. They are routinely prescribed as anti-inflammatory, immunosuppresive, progestational, diuretic, anti-fungal, anabolic and contraceptive agents [1]. Certain steroids have also been used to treat osteoporosis and some forms of breast and prostate cancer, to inhibit HIV-integrase, as well as treat HIV and AIDS [5, 6]. Neurosteroids are those synthesized by glial cells that are present in high concentrations in the central and peripheral nervous system. In a therapeutic context, certain neurosteroids are useful as memory-enhancers, anti-convulsants, and anti-depressants. They have also been shown to have neuroprotective effects against oxidative stress-induced damage, as well as DNA-damage in the brain caused by exposure to mutagens [7]. There is currently an on-going effort to discover new, therapeutically useful steroid compounds, to add to the extant battery of thousands.  1.1.3  Steroids in the Environment  Steroids are released in large quantities in the environment through excretion and decomposition of eukaryotes, as well as in waste from industrial processes [8]. The fate of hormone steroids is potentially the best studied due to the concern for endocrine disruption in  2  humans, fish and other wildlife. Hormone steroids are released into the environment through human sewage and wastewater treatment, as well as animal waste disposal [9]. Cattle are an especially large source of livestock-generated hormones as they are often given steroids to control ovulation and induce abortion, or fed muscle-building androgens to promote growth [10]. Human females release large quantities of estrogenic steroids when menstruating, or especially while pregnant. Estrogen-like steroids also get released through the urine of females taking contraceptives [11]. The presence of estrogenic steroids is especially problematic in aquatic environments. They have been detected in ground water, surface water, and one study even found an average concentration of 0.7 ng/L of estrogenic steroids in drinking water in the south of Germany [2]. Estrogenic steroid contamination can interrupt egg protein production, and can result in the feminization of freshwater fish, higher levels of phytoestrogens in agricultural crops, as well as harmful effects on the reproductive systems of humans [12, 13]. Aquatic systems downstream of pulp and paper mill effluent have exhibited the masculinization of female fish, indicating the presence of androgenic steroids [14]. The treatment of sewage and industrial waste requires a major improvement to prevent this unfettered release of steroids into the environment. The application of biotechnology is a potentially effective and ecofriendly method to deal with steroid contamination.  3  1.2  Microbial Steroid Degradation  Microorganisms have evolved diverse strategies to uptake, transform and degrade steroid compounds. In the natural environment, microbial steroid degradation is an important part of global carbon cycling. In aerobic environments, microorganisms transform steroids released by eukaryotic excretion or decay into biomass and carbon dioxide [4]. Microorganisms within the human body have also evolved to partly metabolize steroids encountered in the digestive system [3].  1.2.1  Harnessing Microbial Steroid Catabolic Pathways for Industrial Purposes  Highly regio- and stereo-selective reactions are required to produce therapeutically and commercially valuable steroids. These reactions often require extreme conditions, temperatures, and hazardous reagents in order to synthetically produce the desired steroid. The expense and energy requirement of synthetic steroid production has led to the development of some high-yield biological production of steroid therapeutics that requires mild reaction conditions and can be performed without the hazardous reagents required in chemical synthesis. Because biological processes are less energetically costly, they are not only more environmentally friendly but also cheaper than chemical syntheses [15]. Such biotechnology essentially converts a readily available starter steroid into a commercially desirable one by using microorganisms to perform some or all of the transformation reactions. Because of the processing required to isolate, purify, and stabilize enzymes and to regenerate their intermediates, it is more economical to use whole cells for  4  steroid manufacture [16]. Cholesterol and β-sitosterol (Figure 1.2) are most often used as the raw materials for microbial transformation into C17-ketosteroids, which are then used as the precursors for the synthesis of various steroid drugs [17]. Cholesterol is mainly extracted from animal sources like wool grease, lard or milk fat, yet despite its good properties for microbial transformation, it is more expensive and less widely-available than β-sitosterol, which is extracted from plants [18]. In order to find cheaper raw materials, there is a great interest in using waste products from pulp and paper mills, or vegetable oil production, as sources for starter steroids. For example, Dias, et al. [19] reported the use of steroids harvested from the above waste products as starting materials for transformation by Mycobacterium sp NRRL 3805 into androstadienedione, which can then be made into contraceptives and corticosteroids.  -sitosterol  Cholesterol  Figure 1.2 -Sitosterol and cholesterol; the two most common starter steroids for whole cell bioconversion [15].  Research in the field of steroid biocatalysis began around 1950 with the discovery of a Rhizopus species capable of performing a crucial step in the synthesis of steroids with useful biological activity [1]. Since then, microbial bioconversion of steroids has been 5  implemented in the large scale production of natural or modified steroid analogues, focusing mainly on steroid hydroxylation, ∆1-dehydrogenation, and sterol side-chain cleavage [15]. Production of steroid drugs is one of the most successful applications of microbial technology in large scale industrial processes. However, the power of microbial steroid transformation is not being harnessed to its full extent – only a fraction of the microbial genes and enzymes capable of transforming steroids are well characterized. A more thorough understanding of how bacteria take up and metabolize these compounds is required in order to better exploit microbes in the manufacture of steroids.  1.2.2  Steroid Bioremediation using Microorganisms  Steroid metabolism is also an important aspect of bioremediation. One study by Chevron Petroleum Technology Company examined the effects of landfarming on waste petroleum sludge at Chevron’s refinery in Perth Amboy, New Jersey [20]. Landfarming involves mixing contaminated soil with uncontaminated soil, and changing conditions, such as moisture content, aeration and pH, to maximize the rate of contaminant degradation. This study used hydrocarbons indigenous to and ubiquitous in crude oils as biomarkers for the progress of bioremediation. Chief among these biomarkers were steroid hydrocarbons including steranes, monoaromatic and triaromatic steroids, and steroids with the cholestane side chain. They found that even in the most degraded samples, biodegradation of these compounds had proceeded only partially to completion. Another study by Jacobsen, et al. [21] showed the necessity for viable soil microorganisms to mineralize steroids in a range of conditions, including temperatures, and  6  soil types. They also noted that the organic amendments brought on by the addition of animal manure altered the pathways and kinetics of steroid dissipation. The effects of animal waste on microbial steroid transformation are relevant in a real-world context, as effluent from sewage treatment plants or agricultural run-off will undoubtedly effect the functioning of the receiving microbial community in soil or water systems. Thus, any bioremediation attempts on animal waste-contaminated sites must take this into account. Oil-contaminated sites, as well as sewage-treatment plant and agricultural effluent, are ideal candidates for “bioaugmentation”, i.e. treating the site with natural or genetically engineered bacteria known to catabolize complex hydrocarbons, especially steroid molecules. With more detailed knowledge of the genes and enzymes bacteria use to transform and degrade steroids, more efficient cocktails of bioremediating strains can be created to specifically deal with different kinds of steroid pollution.  1.3  The Seco-steroid Degradation Pathway  The microbial degradation of cholesterol, testosterone, androstenedione and various bile salts has been the subject of intensive study since the 1950s [15]. In the microorganisms studied so far, it has been shown that the degradation pathways of cholesterol, testosterone and the bile acids merge at the generation of 1,4-androstadiene-3,17-dione (ADD) (Figure 1.3, III) and its respective derivatives, which may have variable side chains at C17 and hydroxyl or ketone groups at other positions on the steroid nucleus. The aerobic degradation of ADDs proceeds via the 9,10-seco-pathway, so called because of the generation of steroids with an opened B-ring, which are referred to as “seco-steroids” [22]. The steps leading up to  7  the generation of ADDs happen differently depending on the starting steroid. The secosteroid pathway results in the formation of 2-hydroxyhexa-2,4-dienoic acid (2-HHD) (Figure 1.3, VIII) from the former A-ring, and an acidic perhydroindane derivative composed of the former B-ring, and rings C and D, called 9,17-dioxo-1,2,3,4,10,19-hexanorandrostan-5-oic acid (DOHNAA) (Figure 1.3, IX). The further conversion of DOHNAA is still under investigation [23, 24]. Details on what has been elucidated for each steroid’s catabolic pathway are discussed separately and in detail in the sections to follow.  1.3.1  Microbial Testosterone Degradation  Comamonas testosteroni is well known for its ability to transform testosterone. The intermediate compounds it generates during testosterone degradation were identified in the 1960s. Guided by these intermediates, the main pathway of testosterone degradation in C. testosteroni was predicted (Figure 1.3), and the genes involved in the pathway have been the focus of intensive study ever since [25].  8  Figure 1.3 Proposed testosterone degradation pathway of C. testosteroni TA441. Testosterone, (I); 4androstenedione (II); androsta-1,4-diene-3,17-dione (III); 9-hydroxyandrosta-1,4-diene-3,17-dione (IV); 3-hydroxy-9,10-secoandrosta-1,3,5(10)-triene-9,17-dione (V); 3,4-dihydroxy-9,10-secoandrosta-1,3,5(10)triene-9,17-dione (VI); 4,5-9,10-diseco-3-hydroxy-5,9,17-trioxoandrosta-1(10),2-diene-4-oic acid (VII); 2hydroxyhexa-2,4-dienoic acid (VIII); 9,17-dioxo-1,2,3,4,10,19-hexanorandrostan-5-oic acid (IX); 9,17dioxo-1,2,3,4,10,19-hexanorandrostan-5-oic acid CoA ester (X) [24]. Multiple arrows indicate more than one enzymatic reaction. Enzymes listed are discussed in detail in the text to follow.  9  1.3.1.1  17-Dehydrogenation  The testosterone pathway begins with the initial 17β-dehydrogenation of the hydroxyl group at C17. 17β-Dehydrogenase (17β-HSD) was isolated from C. testosteroni TA441 and found to act on both the 3β-hydroxyl and 17β-hydroxyl groups of steroids [26]. A mutant strain of TA441 with a disrupted 17β-HSD gene did not show significant growth on epiandrosterone, which has a 3β-hydroxyl group and a 17-ketone group. However, with noticeably slower growth than wild type, it did degrade testosterone, which has a 17βhydroxyl and a 3-ketone group (I). Dehydrogenation at the C17 position is thought to be a reversible reaction, as the intermediate compounds isolated with a 17-ketone group are often accompanied by a small amount of analogs with a C17 hydroxyl group. Thus, C. testosteroni likely has more than one enzyme capable of 17β-dehydrogenating/hydrogenating activity, while it requires 17β-HSD to degrade 3β-dehydroxy steroids [24]. Therefore, this enzyme might more logically be referred to as 3β-HSD.  1.3.1.2  Transformation of Androstenedione  The dehydrogenation of the 17β-hydroxyl group of testosterone produces 4androstene-3,17-dione (4-AD) (II). In microbial steroid degradation, a ketone group at C3 and the double bonds, 1 and 4, are essential for cleavage of the B-ring and aromatization of the A-ring. 1-Dehydrogenase (TesH) converts 4-AD to ADD (III) with the insertion of the 1 double bond. Subsequently, the oxygenase, 9α-hydroxylase, forms an unstable  10  intermediate (IV), leading to spontaneous cleavage of ring B between C9 and C10 and aromatization of the A-ring, producing 3-hydroxy-9,10-secoandrosta-1,3,5(10)-triene-9,17dione (3-HSA) (V). ORF17 is proposed to encode the reductase component of 9αhydroxylase, because an ORF17-disrupted mutant of strain TA441 accumulates ADD when incubated with testosterone [27]. TesA1A2 hydroxylates 3-HSA at C4 to produce 3,4-dihydroxy-9,10-secoandrosta1,3,5(10)triene-9,17-dione (3,4-HSA) (VI). 3,4-HSA is then cleaved between C4 and C5 by meta-cleavage enzyme TesB to produce 4,5-9,10-diseco-3-hydroxy-5,9,17-trioxoandorosta1(10),2-diene-4-oic acid (4,9-DHSA) (VII). The hydrolase TesD then hydrolyzes 4,9-DHSA to produce 2-HHD (VIII) and DOHNAA (IX). TesEGF degrade what is left of the A-ring in a process similar to the bacterial degradation of aromatic compounds [24]. ORF18 was shown to be necessary for the further degradation of DOHNAA, as ORF18-disrupted mutants accumulate DOHNAA when incubated with testosterone. Based on database searches, ORF18 was implicated in encoding a CoA-synthetase, suggesting the degradation of DOHNAA is initiated by the addition of CoA (X) [28].  1.3.1.3  Transformation of DOHNAA  Further degradation of DOHNAA has not yet been elucidated, however analysis of intermediates accumulated by gene-disrupted mutants involving the ORF1-33 of strain TA441 suggest degradation is initiated mainly via β-oxidation [24].  11  1.3.1.4  Steroid Pathway Gene Clusters in C. testosteroni  Genes for the degradation of steroid structures make up two major gene clusters in C. testosteroni. The first is composed mainly of genes involved in the aromatization and cleavage of the A-ring (named tesG to ORF18 in strain TA441) and the second consists of genes involved in the β-oxidation of the B,C,D-rings (named steA to tesR in strain TA441). Recently these two gene clusters were shown to be several dozens of kilobases apart on the same DNA strand, with quite a few possible steroid degradation genes located in the intervening DNA, suggesting this DNA region of more than 100 kb may be a large steroid degradation gene locus of C. testosteroni [29].  1.3.2  Microbial Cholesterol Degradation  Microbial transformation of steroids has recently also received attention from biomedical researchers due to the hypothesis that Mycobacterium tuberculosis, the leading cause of mortality from bacterial infection, may use host cholesterol during lung infection [30].  1.3.2.1  Modeling M. tuberculosis Cholesterol Degradation in R. jostii RHA1  Recent genomic analysis showed that approximately 60% of the 3,999 genes of M. tuberculosis are conserved in the related Actinobacterium Rhodococcus jostii RHA1, including many genes of unknown function. Because of this genetic similarity, RHA1 has  12  been used as model organism for the cholesterol catabolic pathway of M. tuberculosis, in hopes of shedding light on how M. tuberculosis utilizes host cholesterol during infection [23].  1.3.2.2  Cholesterol Uptake  The genome sequence of M. tuberculosis revealed four sets of “mammalian cell entry” genes (mce), mce1-4, which are of particular interest due to their predicted role in pathogenesis [31]. Deletion of nearly the entire mce3 or mce4 operon increased long term survival of mice after low dose aerosol infection, and a decreased bacterial burden was also observed in infections with the mce4 deletion mutants [32]. Based on bioinformatic analysis, it is highly likely that the mce1 and mce4 loci encode novel types of ABC transporters. It has been demonstrated that the Mce4 system is involved in steroid transport, and is specifically required for the uptake of cholesterol, 5-α-cholestanol, 5-α-cholestanone, and β-sitosterol; all steroids with similar structures [33]. Thus the cholesterol catabolic pathway of Rhodococci and Mycobacteria requires the Mce4 system to transport the steroid into the cell.  1.3.2.3  Cholesterol Side-Chain Degradation and A-ring Oxidation  The microbial degradation of cholesterol involves two processes: side-chain degradation and the degradation of the rings of the steroid nucleus. The order of these events may vary depending on the organism [34]. In RHA1 it was found that oxidation of the side chain precedes oxidation of the 3β-hydroxyl moeity of the A-ring. CYP125 is a steroid 26-  13  monooxygenase that catalyzes the formation of C26 to the corresponding carboxylic acid, or to an intermediate, to initiate degradation of the side chain (Figure 4) [35, 36]. Further degradation of the side chain proceeds in a process similar to the β-oxidation of fatty acids. Several acyl coenzyme A (CoA)-ligases, dehydrogenases, hydratases and thiolases are thought to catalyze the activation and thiolytic cleavage of the cholesterol side chain. This process releases two propionyl-CoA plus one acetyl-CoA [37]. RHA1 has a series of genes that are upregulated during growth on cholesterol that encode the types of enzymes necessary for this process [23].  Figure 1.4 Proposed cholesterol (I) side-chain degradation pathway. Double arrows indicate more than one enzymatic reaction.  Depending on the strain, either an O2-dependent cholesterol oxidase (CHO) or 3βhydroxysteroid dehydrogenase (3β-HSD) catalyzes the generation of 4-cholestene-3-one by oxidating the 3β-hydroxyl moeity and isomerizing 5 into 4 (Figure 1.5) [35]. Accumulation 14  of 4-AD (II) and ADD (III) has been detected in some mutants of Mycobacterium sp. suggesting that further side-chain degradation occurs before appreciable degradation of the steroid rings. However, in certain RHA1 mutants, accumulations of 3-HSA with a C17 isopropyl side chain were detected, thus in certain strains, the order of the degradation of the side chain and rings may vary [37].  Figure 1.5 Deduced cholesterol nucleus degradation pathway. Cholesterol, (I); AD (II); ADD (III); 9OHADD (IV); 3-HSA (V); 3,4-DHSA (VI); 4,9-DHSA (VII); DOHNAA (VIII); 2-HHD (IX). Dashed arrows indicate multiple enzymatic steps. The C17 side-chain is represented as R as the progress of sidechain degradation with regards to ring oxidation may vary depending on the strain.  15  1.3.2.4  Predicting Cholesterol Pathway Gene Function  A recent study by van der Geize, et al. [23] found that during growth of RHA1 on 2 mM cholesterol, 572 genes were variously upregulated when compared to growth on pyruvate. A cluster of 51 genes was found to encode proteins that have significant sequence identity with already-characterized enzymes involved in the catabolism of rings A and B in strain TA441 and Rhodococcus erythropolis SQ1, such that the function of several of these RHA1 enzymes can be confidently predicted.  1.3.2.5  Rings A and B Degradation and DOHNAA Generation  KSTD, a 3-ketosteroid 1-dehydrogenase, performs the 1-dehydrogenation of C1, generating ADD or a derivative with a partial side-chain (III) [38]. KshAB, encoding a two component 3-ketosteroid 9α-monooxygenase, introduces the 9α-hydroxyl moeity into the Bring, generating 9α-hydroxy-4-androstene-3,17-dione (9OHADD) (IV) [39]. This allows for the nonenzymatic hydrolysis of 9OHADD into 3-HSA (V). The enzymes KSTD, KshA and KshB of RHA1 are 40-69% identical to their orthologs identified in R. erythropolis SQ1 [23]. Further degradation of 3-HSA by RHA1 is thought to proceed in the same manner as observed in testosterone degradation by strain TA441. Seven genes, with names beginning with hsa are clustered with kstD and kshA on the RHA1 genome, encoding proteins that share 30-60% amino acid sequence similarity with orthologs encoded by tes genes of strain TA441 [27, 40]. HsaAB, a 2-component, flavin-dependent monooxygenase, hydroxylates ring A of 3-HSA at C4 yielding 3,4-DHSA (VI). HsaC, an extradiol dioxygenase, cleaves 3,4-DHSA 16  to produce 4,9-DHSA (VII). HsaD then hydrolyzes the C5-C6 bond to produce DOHNAA (VIII) and 2-HHD (IX). 2-HHD is further transformed by HsaEFG, a hydratase, an aldolase and an acetaldehyde dehydrogenase respectively, to propionyl-CoA and pyruvate. As in the testosterone pathway of C. testosteroni, the further degradation of DOHNAA in RHA1 and M. tuberculosis is still being investigated [37].  1.3.3  Microbial Bile Salt Degradation  Bile acids are surface active steroids that aid in the digestion of dietary fats in vertebrates [41]. They are highly water-soluble, due to the α-hydroxyl groups at C3, C7 and C12, as well as the five-carbon acyl side chain attached to C17 (Figure 6). Because all the hydroxyl groups protrude in the same direction, bile acids are amphiphilic and act as detergents. As such, bile acids can perturb the structure of the bacterial cell membrane and induce lysis [42].  Figure 1.6 Structure of cholic acid.  17  Bile salts are bile acids with a cation, usually sodium, on the hydroxyl group of the C17 side chain. The ability to degrade bile salts is widespread amongst bacteria, and is wellstudied in anaerobic bacteria of the intestines [4]. Enteric bacteria have evolved diverse physiological adaptations to deal with high concentrations of bile salt: in the duodenum, bacteria face bile salt concentrations of up to 20 mM [43]. Intestinal bile salt transformation is mainly achieved by anaerobic bacteria of the genera Bacteroides, Eubacterium and Clostridium. The benefit of bile salt transformation for intestinal bacteria seems to be the acquisition of glycine and taurine from conjugated bile salts [43, 44].  1.3.3.1  Bile Salt Degradation by Free-Living Bacteria  Considerable quantities of bile salts are released into the environment through fecal matter (300 – 600 mg per day per human) and urine (approximately 4 mg per day per human) [22]. Thus, it is not surprising that many aerobic soil and aquatic bacteria can grow with bile salts as a carbon and energy source. Complete aerobic degradation of bile salts has been reported by various groups of bacteria from Actinobacteria, Betaproteobacteria and Gammaproteobacteria. The best studied bile salt degrading Actinobacteria are members of the genera, Arthrobacter, Corynebacterium, Mycobacterium, Nocardia, Rhodococcus, and Streptomyces [16]. Of the Betaproteobacteria, C. testosteroni is being intensively studied, and of the Gammaproteobacteria, different Pseudomonas species are of interest [4, 24]. For free-living bacteria, bile salts are both a nutrient source and a potential stress factor. For Pseudomonas sp. strain Chol1, it was shown that seco-steroids produced in the degradation pathway have toxic effects. Thus, these microorganisms have likely evolved 18  adaptive strategies to reduce their exposure to toxic bile acid degradation products. One possibility is that the degradation pathway proceeds in such a manner so as to reduce the cell’s exposure to toxic substrates. Several studies support this hypothesis by showing the transient extracellular accumulation of degradation intermediates, which would keep intracellular levels of these toxic compounds low [45]. It is not yet known whether accumulation of these metabolites is the product of diffusion or of active efflux, however efflux systems for non-charged steroids have been found, and a bile salt exporter has been postulated for intestinal bacteria [43]. Transient extracellular accumulation of bile salt degradation products also implies uptake of these intermediates for further conversion, but currently no intermediate uptake systems have been characterized [45].  1.3.3.2  Relevance of Microbial Bile Salt Transformation  For energy conservation, free-living bacteria generally appear to oxidize bile salts completely to CO2, thus bile salt degradation is relevant as a source of atmospheric CO2 [4]. Bacterial bile salt transformation also has applications in biotechnology, as bile salts and their precursors are processed to their desired conformation by biotransformation to be used as treatment for gallstones [46]. Another goal is to develop probiotic strains for introduction to the intestines that transform certain conformations of bile salts, thereby lowering the concentration of secondary bile salts that are presumptive colon cancer promoters [43]. Finally, the pathway for bile salt degradation merges with that of other steroid substrates, such that knowledge of bile salt metabolism is relevant for metabolic engineering of steroidtransforming bacteria in general [4].  19  1.3.3.3  The Unifying Scheme of Microbial Bile Salt Degradation  In 1982, Hayakawa proposed a unifying scheme for bacterial bile salt degradation based on investigations from the 1970s and 1980s that detected intermediates in bacterial cultures growing on bile salts [22]. However, most of what is known about the actual pathway is based on degradation pathways for other steroids. The differentiation of the degradation pathway of cholic acid, the primary mammalian bile salt, from that of testosterone or cholesterol is the processing of the 3α-,7α-, and 12α-hydroxyl groups and the acyl-side chain.  1.3.3.4  Initiation of the Bile Salt Pathway  As with other steroids, the hydroxyl group at C3 must be oxidized to a ketone before the desaturation of the A-ring can occur via double bond insertion between C1 - C2 and C4 C5. Because the C3 hydroxyl group is in the α-orientation as opposed to , as seen in cholesterol and testosterone, it potentially requires a different HSD than the aforementioned steroids. Degradation of the side-chain begins with activation of the acyl-group by Coenzyme A (CoA; Figure 1.7). Formation of cholyl-CoA was detected in strain Chol1, as well as formation of CoA-esters of 3-ketocholate and 1,4-3-ketocholate. Thus it is still unclear whether CoA activation happens before or after A-ring oxidation; like the cholesterol degradation pathway, the order in which these events occur likely varies from strain to strain, or possibly happens simultaneously [4].  20  Figure 1.7 Proposed -oxidation of the cholic acid side chain [45].  1.3.3.5  -Oxidation of Bile Salts in Pseudomonas sp. Strain Chol1  Birkenmaier, et al. [45] did the first genetic and biochemical study on the β-oxidation of bile salts. They created the R1 transposon mutant of strain Chol1, with a disrupted acad gene, that accumulates the compound 7α,12α-dihydroxy-3-oxopregna-1,4-diene-20carboxylate (DHOPDC; Figure 1.8), which is similar to AD but has a 7α-hydroxyl and a 12αhydroxyl group, and an isopropionyl side chain. The acad gene encodes a putative acyl-CoAdehydrogenase, which inserts a double bond into the side chain of DHOPDC-CoA. Accumulation of DHOPDC has also been detected in a mutant of strain TA441 with a  21  transposon inserted in a gene with 70% similarity to strain Chol1’s acad [45]. The compounds that have been identified as intermediates in cholic acid degradation support the proposed pathway that proceeds via β-oxidation of the side chain [4].  Figure 1.8 Structure of DHOPDC [45].  1.3.3.6  Degradation of Rings A and B  β-Oxidation of the side chain leads to the formation of 7,12-dihydroxy-1,4androsteine-3,17-dione, or DHADD (Figure 1.9).  Figure 1.9 Structure of DHADD [45].  22  In strain TA441, it is postulated that the 12α-hydroxyl group sterically hinders 9αhydroxylase from binding C9, thus it must first be converted to the β-orientation so a 9αhydroxyl can be added to the B ring [24]. Then, opening of ring B and the aromatization of ring A results in the formation of 3,7,12-trihydroxy-9,10-secoandrosta-1,3,5(10)triene-9,17dione (THSATD) (Figure 1.10).  Figure 1.10 Structure of THSATD [45].  During incubations of strain Chol1 with cholic acid, it was found that in the first half of exponential grown, all of the cholic acid was converted to THSATD, which accumulated in the medium and was then degraded in the second half of exponential growth [45]. The rest of the unifying scheme for cholic acid degradation follows the seco-steroid degradation pathway that is observed in testosterone and cholesterol, complete with generation of 2-HHD and 7,12-dihydroxy DOHNAA [4], with the exception that the hydroxyl groups at C7 and C12 are not modified during the above degradation process. In order to shed light on the unique intricacies of microbial cholic acid degradation and how it differs from that of testosterone or cholesterol, detailed studies of the genes and  23  enzymes involved in the pathway are required. In addition, understanding the mechanisms behind the observed efflux and uptake of intermediates can be crucial in a biotechnological context.  1.4  1.4.1  RHA1: An Excellent System to Study Cholic Acid Catabolism  The Unique Properties of Actinobacteria  The Actinobacteria are a group of high-GC, gram-positive bacteria that are ubiquitous in diverse environments. In soil, Actinobacteria provide essential ecosystem services, decomposing and recycling numerous organic compounds [47]. Potentially the most important features of Actinobacteria in determining their ecological niches and fitness are their catabolic versatility and ability to withstand environmental stresses. These capacities at least partially explain the large genomes typical of soil Actinobacteria, as exemplified in RHA1. This soil Actinomycete boasts a large, fully-sequenced 9.7-Mb genome, packed with genes that allow it to catabolize a wide range of aromatic compounds, including carbohydrates, nitriles, steroids, and xenobiotic compounds as sole sources of carbon and energy [48].  1.4.2  The Resilience of Rhodococci  The cell wall of rhodococci is composed of mycolic acids, similar to those found in Mycobacterium. Mycolic acids contain long aliphatic chains that facilitate the uptake and  24  subsequent metabolism of hydrophobic substrates across the cell envelope [49]. Rhodococci can survive in starvation conditions and in many cases, the degradation of pollutants is not repressed in the presence of more easily degradable carbon sources [50]. These attributes make rhodocci highly suitable for environmental remediation. Because they grow rapidly and robustly on a wide variety of substrates, they are highly useful in biotechnological applications in the pharmaceutical and chemical industries, primarily for biotransformation and biodegradation of organic compounds [51]. Likewise, they are good candidates for the industrial production of bioactive steroid compounds, but we must first understand more about the mechanisms rhodococci use to transform steroids [15].  1.4.3  Bile Salt Pathway Genes in RHA1  RHA1 is an excellent system to study microbial steroid catabolism. The genome of RHA1 appears to encode at least four distinct steroid catabolic pathways. Microarray analysis of this organism grown separately on cholesterol, androstenedione, and cholic acid revealed over 700 genes that were variously upregulated on these three steroids (Mohn, W.W., et al. unpublished). Guided by this transcriptomic analysis, research focused on cholesterol degradation has identified several genes in RHA1 involved in cholesterol uptake, side-chain removal, and initial ring degradation [33, 48]. A homologous yet distinct degradation pathway for cholic acid catabolism is also implicated by the transcriptomic data. Several genes are upregulated that are likely involved in the degradation of the cholic acid side chain, and rings A and B. In addition to pathway genes, genes encoding an ATP-binding cassette (ABC) transporter (ro04888 – ro04885, Figure 1.11) and a multi-drug transporter of  25  the MFS superfamily (ro05792, Figure 1.12) are also highly upregulated by RHA1 growing on cholic acid. The ABC transporter appears to include an individual permease and ATPbinding components as well as a fused permease/ATP binding component.  Figure 1.11 ATP-binding cassette transporter.  Figure 1.12 Multi-drug transporter of the MFS superfamily: A. outward-facing conformation, B. inwardfacing conformation.  26  1.4.4  The Aim of this Study  This study examined two transporters associated with cholic acid catabolism. Knockout mutants of RHA1 were generated in which the above ABC and MFS transporters were individually inactivated using unmarked, in-frame gene deletion. Growth of the wildtype and mutant strains on cholic acid was characterized, and metabolite production was determined. Like other organisms grown on cholic acid, RHA1 also exhibits the transient accumulation of extracellular metabolites, which are reassimilated and further degraded in later exponential and stationary growth phase. Neither transporter was required for uptake of cholic acid, but each was essential for the assimilation of particular metabolites, which were structurally elucidated. To our knowledge, this is the first study to identify and characterize transporters required for the assimilation of excreted steroid metabolites. Preliminary structural analysis suggests these extracellular metabolites are very structurally similar to those that accumulate in studies of other organisms grown on cholic acid [4, 45], except that the THSATDderivative of RHA1 retains an isopropyl-side chain at C17. This implies that RHA1 also degrades cholic acid via the same pathway as previously characterized gram-negative bacteria, but that RHA1 differs with respect to the order of nucleus and side chain degradation. This study contributes to the knowledge of the bacterial physiology behind certain steroid transformations, as well as demonstrates that during microbial cholic acid catabolism, A-ring oxidation, cleavage of the B-ring, and potentially even generation of 7,12-dihydroxy-  27  DOHNAA, can precede complete degradation of the C17 side chain in some strains. In addition, this study sheds new light on the diversity of functions that transport proteins can perform. Finally, this research contributes to the fundamental knowledge of the physiology, genetics, and ecology of the industrially and environmentally important members of Rhodococcus and related genera.  28  Chapter 2: Materials and Methods  2.1  Reagents and Chemicals  All reagents and chemicals were ordered from Fisher Scientific, Sigma, Bio Rad, Thermo Scientific, and Invitrogen and used without further purification unless specified. Cholic acid was at minimum 98% pure. Restriction Enzymes for cloning were purchased from New England Biolabs. T4 DNA ligase was purchased from Fermentas. Water for media preparation and experimental procedures was purified to a resistivity greater than 10 MΩ·cm using a Thorton 200CR Conductivity/Resistivity Instrument from Mettler Toledo.  2.2  Cell Growth Assays  RHA1 and derivative strains were cultivated at 30°C and 200 rpm in M9 mineral medium [52] supplemented with Goodies [53]. This is composed by creating a stock salt solution (Table 2.1), adding 50 ml of the stock salts to 100 ml total of a concentrated Goodies solution (Table 2.2), adding 75 ml of this concentrated Goodies solution to 300 ml total of a Mg/Ca/B1/Goodies mix (Table 2.3) and finally combining 1 ml of this Goodies mix with 10 ml of the M9 basal salts solution (Table 2.4), and bringing the volume to 100 mL with H2O. For a carbon source, either 2.0 mM steroid substrate or 20 mM pyruvate was added. Escherichia coli strains were grown at 37°C in Luria-Bertani (LB) broth. Bacto agar (1.5% w/v) from Difco was used for solid medium. Culture growth was measured as the  29  change in optical density at 600 nm. Biomass measurements, based on protein measurements, were performed with a BCA protein assay from Thermo, Scientific, using bovine serum albumin as the standard. To test for transformation of cholic acid metabolites, M9 mineral medium was diluted one-fold with filter-sterilized strain camB or camM culture supernatant harvested during early stationary growth phase.  Table 2.1 Stock salt solution recipe.  Chemical MgCl2-6H2O CaCO3 FeSO4-7H2O ZnSO4-7H2O MnSO4-H2O CuSO4-5H2O CoCl2-6H2O H3BO3 HCl  g/L 22.94 2.0 4.5 1.44 0.85 0.25 0.24 0.06 51.3 mL  Table 2.2 Concentrated goodies recipe (100 mL).  Component Stock Salt Solution MgSO4 1% FeSO4 H2O  Quantity 50 mL 3.009 g 25 mL Adjust volume to 100 mL  30  Table 2.3 100X Mg/Ca/B1/Goodies mix recipe (300 mL).  Component MgSO4 CaCl2 10 mM thiamine (vitamin B1) Concentrated Goodies H2O  Quantity 60 mL 3 mL 3 mL 75 mL Adjust volume to 300 mL  Table 2.4 M9 basal salts recipe for 1 L.  Chemical KH2PO4 Na2HPO4 NaCl NH4Cl  2.3  Quantity 30 g 60 g 5g 10 g  DNA Manipulation and Construct Preparation  DNA was amplified, purified, and manipulated according to standard procedures [52]. Polymerase chain reactions (PCR) were performed with Taq Polymerase from Qiagen or High-Fidelity Expand Polymerase from Roche in a 96-well PTC-200 Peltier Thermal Cycler. Oligonucleotides were designed with GeneRunner software and synthesized at the Nucleic Acid Protein Services Unit at UBC (NAPS). Nucleotide sequencing of clones and constructs were also performed at NAPS. The mutant strains, camB and camM, were generated, based on previously described methods, by amplifying 800 – 1000 bp fragments that flank the gene of interest, with less than 20% of the targeted gene included in the amplicons [38]. The camB and camM upstream flanking regions were amplified from fosmid DNA (RF00110B17 and 31  RF00110H09, respectively; Table 2.5) with the following primers: cBupFor, cBupRev, cMupFor and cMupRev (Table 2.6). The downstream flanking regions were amplified in a similar fashion using cBdwFor, cBdwRev, cMdwFor, cMdwRev primers (Table 2.6). Tripleligations using T4 DNA ligase, with the cut sites for EcoR1, HindIII and SphI for camB and EcoRI, HindIII and XbaI for camM, were performed to clone the amplicons into pK18mobSacB (Table 2.5) to produce pK18camB and pK18camM respectively. The mutagenic constructs were transformed by electroporation and propagated in DH5α E. coli, chosen for its high transformation efficiency. Time constants of 4.5 volts or higher were achieved for all transformations via electroporation.  Table 2.5 Fosmids and plasmids used in this study.  Relevant characteristicsa Reference/origin r Amp [48] r Amp [48] CmrKanr [54] CmrKanr, pk18mobSacB camB deletion plasmid This study r r Cm Kan , pk18mobSacB camM deletion plasmid This study r -C31 Am , int [55] Amr, int-C31, pSET152 camB complementation This study plasmid pSTcamM Amr, intphi-C31, pSET152 camM complementation This study plasmid pTIP AmprCmr [56] r r pTIPcamB Amp Cm , pTIP camB complementation plasmid This study a Ampr: ampicillin resistance, Cmr: chloramphenicol resistance, Kanr: kanamycin resistance, Amr: apramycin resistance, int-C31: bacteriophage -C31 attachment site. Fosmids/Plasmids RF00110B17 RF00110H09 pK18mobSacB pK18camB pK18camM pSET152 pSTcamB  For the complementation assay, primers were designed to include in the amplicons at least 200 base pairs upstream of the target gene or target operon, thereby including the native promoter region. The camB and camM amplicons for the complementation assay were  32  produced from fosmid DNA (RF00110B17 and RF00110H09, respectively; Table 2.5), using primers camBpSTf and camBpSTr for camB and camMpSTf and camMpSTr for camM (Table 2.6). DH5α transformants were screened using colony PCR. Colonies were sampled with a toothpick, suspended in 12.5 l of water with 50 g/ml proteinase K, incubated at 50C for 30 minutes and 80C for 20 minutes, and then centrifuged at 16,000 x g for 5 minutes to remove cell debris. Colony PCR reactions contained 2 l of the above cell extract, in a total volume of 10 l. Successful DH5α transformants were grown for 16 hours at 37C in LB, and their plasmid constructs were then isolated using the Bio Rad plasmid miniprep kit for subsequent transformations.  33  Table 2.6 Oligonucleotides used in this study.  Primer name Sequence (5’ to 3’)a cBupFor GCGAAGCTTCGAGATGAAGACGATGTTGTAGCG cBupRev CGGGAATTCGCGCAGAACAAGATCCTCTCGTTC cBdwFor CGGAAGCTTGCGGTCTTCGTGGTCCTCATC cBdwRev GCGGCATGCACCCACCGAGGAGGCCAGTACC cMupFor TCTGAATTCGGTAGTGGACGAGGGTGGCGAC cMupRev TTATCTAGAGGGATTCGAGAGGATCAGCACG cMdwFor TGTTCTAGACTCCTCATCCGCAACAAGATCGAG cMdwRev TCCAAGCTTCTTTCGACGTGGTCCTCAGCG camBColF GACGACCAGCGCGAGGATGCAG camBColR GTCGAAACCCGGAGACGAGGCGTAC camMColF ACCAGTAGGGCGCGATCTGATGGAG camMColR AGGTGACCCGGCGCGTCAATC camBpSTf ATATCTAGAAGACACGTTCGCCGAGTCGGTC camBpSTr AATTCTAGAAGATGTTCGCCAAGACCATCG camMpSTf AAAGAATTCAGACTGAGTGATTCCGTCATGGG camMpSTr AAATCTAGATGAACACTTGGACTGCCACACCG camBpTIPf CTTGTTACATATGACCATCGTCTGGTCGGGCCTC camBpTIPr ATCAAGCTTTCACACCACCCGTTCCCGCACC camBQFor TCTCGTCCCACACGTTCAAC camBQRev CGAAGGTTCCGACCATCATC camMQFor CCGTCGCGAAAACATTGG camMQRev GGATCGGTCATCACGAACATC a The indicated restriction enzyme cut sites are underlined.  Restriction Site EcoRI HindIII HindIII SphI EcoRI XbaI Xba1 HindIII --------XbaI XbaI EcoRI XbaI NdeI HindIII ---------  34  The construct pTIPcamB (Table 2.5) was generated by amplifying a wild type copy of camB with the camBpTIPf and camBpTIPr primers (Table 2.6). A double-ligation was used to clone this amplicon into pTIP cut with both NdeI and HindIII, and the pTIPcamB construct was then directly transformed into electrocompetent camB cells via electroporation using previously described methods [57].  2.4  Cloning of Knockout Mutants  Gene deletion mutants camB and camM were generated by the previously described sacB counterselection system [38]. Wild Type RHA1 was cultivated in Lysogeny Broth Peptone media (LBP) containing 30 g/ml nalidixic acid at 30C for 48 hours at 200 rpm. These cultures were then plated on solid LBP agar containing 30 g/ml nalidixic acid and incubated at 30C for an additional 48 hours. The mutagenic constructs pK18camB and pK18camM were transformed by electroporation into E. coli S17-1 donor cells, which were plated on LBP plates containing 25 g/ml kanamycin and grown for 24 hours at 37C, then left at room temperature for an additional 24 hours to garner more cell material. Cells from both the donor strain and the RHA1 plates were suspended in 2 ml of LBP media, and aliquots of 750 l each were gently mixed together. The cells were pelleted by centrifugation (9,400 x g for 1 min), re-suspended in 1 ml of LBP media, and aliquots of 200 l were spread on non-selective LBP plates and incubated overnight at 30C to facilitate conjugation. Cells were then harvested from plates with confluent growth, suspended in 2 ml of LBP media, and aliquots of 100 l were spread on selective LBP plates with 50 µg/ml kanamycin plus 30 µg/ml naladixic acid and incubated at 30C for 96 hours. 35  To initiate the integration of the pK18mobsacB construct into the RHA1 genome, kanamycin-resistant colonies were selected and replica-plated on LBP with 10% (w/v) sucrose and on LBP with 50 g/ml kanamycin, each additionally containing 30 g/ml nalidixic acid. Within two days, colonies that were sucrose sensitive and kanamycin resistant (Kanr/Sucs) were selected and cultured overnight in 25 ml LBP medium at 30C. Aliquots of 25 l were spread on LBP plates supplemented with 10% sucrose, and incubated at 30C for 72 hours. Sucrose resistant colonies were replica-plated on LBP plates as well as LBP supplemented with 50 g/ml kanamycin. Kanamycin sensitive colonies (Kans) were screened by colony PCR using camBColF and camBColR for the camB deletion and camMColF and camMColR for the camM deletion (Table 2.6). The regions of deletion in strains camB and camM were sequenced to ensure there were no changes in the regions flanking the DNAdeletion.  2.5  Cloning of Complementation Strains  The entire 5-kb, four-gene operon camABCD and the 1.8-kb single-gene camM were cloned to generate the complementation strains camBc and camMc. Double-ligations using T4 DNA ligase with the cut sites XbaI for camBc and EcoRI and XbaI for camMc were performed to clone the amplicons into the pSET152 integrative vector, to produce pSTcamB for camB complementation and pSTcamM for camM complementation (Table 2.5). XbaI was used to cut both ends of the camB amplicon because it was the only restriction enzyme in the pSET152 multiple cloning site that was also a non-cutter in the  36  camB operon. The complementation constructs were transformed via electroporation into DH5α E.coli. The mutants camB and camM were individually cultivated in LB media containing 30 g/ml nalidixic acid at 30C for 48 hours at 200 rpm. These cultures were then plated on solid LBagar containing 30 g/ml nalidixic acid and incubated at 30C for an additional 48 hours. The complementation constructs pSTcamB and pSTcamM were transformed by electroporation into E. coli S17-1 donor cells and plated on LB plates containing 50 g/ml apramycin and grown for 24 hours at 37C, then left at room temperature for an additional 24 hours to garner more cell material. Cells from both the donor strain and the RHA1 plates were re-suspended in 2 ml of LB media, and aliquots of 750 l each were gently mixed together. The cells were pelleted by centrifugation (9,400 x g for 1 min), re-suspended in 1 ml of LB media, and aliquots of 200 l were spread on non-selective LB plates and incubated overnight at 30C to facilitate conjugation. Cells were then harvested from plates with confluent growth, suspended in 2 ml of LB media, and aliquots of 100 l were spread on selective LB plates with 50 µg/ml apramycin plus 30 µg/ml nalidixic acid and incubated at 30C for 96 hours to facilitate integration in the -C31 bacteriophage attachment site. Subsequent colonies were re-plated on selective LB plates with 50 µg/ml apramycin plus 30 µg/ml nalidixic acid and incubated at 30C for 48 hours, then screened for the presence of both the truncated and the wild type target gene using cBupFor and cBdwRev primers for camBc and cMupFor and cMdwRev primers for camMc (Table 2.6). The pSET152 vector cannot exist freely in Rhodococcus strains [55].  37  Preliminary attempts were made to complement the camB mutant using the inducible expression vector pTIP as previously described [56]. This approach was soon abandoned due to unreliable results, and the pSET152 method was adopted instead.  2.6  RNA Extraction  Samples of 10 ml were taken when cultures reached approximately 35%, 65% and 100% of their maximal OD600., corresponding to exponential, late exponential and stationary phases, respectively. A 1/10 volume of stop solution (95% reagent alcohol: 5% acid phenol [v/v] , pH 5) was added to samples, which were then centrifuged at 4,000 x g and 4C for 10 minutes. Cells were suspended in 1.0 ml supernatant plus 2.0 ml of RNA Protect from Qiagen, incubated for 5 minutes at room temperature, then centrifuged at 18,000 x g for 2 minutes. Pellets were frozen on dry ice and stored at -80C. RNA extraction and purification were performed based on previous methods [58] with the following modifications. RNA was isolated from cell pellets by vortexing with glass beads in hot phenol plus sodium dodecyl sulfate at final concentrations of 14.3% and 0.9%, respectively. Cell debris was removed with acetate plus phenol-chloroform [1:1, v/v]. Nucleic acids were precipitated with 0.6 M acetate plus 100% isopropanol, and treated with DNase I from Invitrogen, then incubated at 37C for 1 hour with RNase Out from Invitrogen. RNA was extracted and precipitated with phenol-chloroform [1:1, v/v], then purified with the Qiagen RNeasy minikit, according to instructions.  38  2.7  cDNA Synthesis  Synthesis of cDNA was performed with the commercially available ThermoScriptTM RT-PCR System protocol from Invitrogen. All reagents were provided from Invitrogen unless otherwise indicated. 5 g RNA were incubated with 50 ng of random hexamers and 10 mM dNTP Mix. The mixture was brought to 12 l with diethyl pyrocarbonate (DEPC)treated water, and incubated at 65C for 5 minutes, then cooled on ice for 10 minutes. The following mix was prepared and added to each reaction: 5X cDNA Synthesis Buffer, 0.1 M DTT, 10 U RNaseOUTTM, 15 U ThermoScriptTM RT, brought to 10 l with DEPC-treated water. The samples were incubated at 25C for 10 minutes, 50C for 50 minutes, then 85C for 5 minutes. 2 U of RNase H was added and the samples were incubated at 37C for 20 minutes. Fresh cDNA samples were used immediately for RT-QPCR analysis.  2.8  RT-QPCR Analysis  Reverse-transcriptase quantitative PCR (RT-QPCR) measured the expression of targeted genes compared to that of an internal standard gene, ro01702, encoding a DNA polymerase IV. Previous microarray experiments showed that this housekeeping gene is expressed as an approximately constant fraction of total mRNA under a variety of growth conditions and during all growth phases [58]. The TaqMan probes and primers used in this study were designed with the Primer Express Software v2.0 from Applied Biosystems. Each QPCR reaction mixture contained 1.0 l of the 20 l of cDNA produced, 200 nM TaqMan probe, and 10 l 2X TaqMan universal PCR master mix from Applied Biosystems, in a total 39  volume of 20 l. Duplicate singleplex reactions for the housekeeping gene and the target gene were performed in a Stratagene Mx3000P real-time PCR system for 2 minutes at 50C, 10 minutes at 95C and then 40 cycles of 15 seconds at 95C and 1 minute at 60C. The quantitative PCR cycle threshold (CT) results were analyzed using the comparative CT method (CT method). The Student’s one-sample t test was used to evaluate whether the gene targets were differentially regulated. The expression ratios were calculated with the formula E-CT, where E is the average amplification efficiency derived from the standard curves for the target and housekeeping genes [59].  2.9  Chemical Analysis  For gas chromatography mass spectrometry (GCMS) analysis, samples were periodically collected and cells were removed by centrifugation. The supernatant was then acidified with a 1/50 volume 10% H2SO4, extracted twice by mixing at room temperature for 30 minutes with an equal volume of ethyl acetate, dried under nitrogen and dissolved in 200 l pyridine. An internal standard of 0.25 mM cholestane was used to normalize each sample. The extracts were derivatized at room temperature with 200 l bis(trimethylsilyl)trifluoroacetamide/trimethylchlorosilane (99:1) from Sigma-Aldrich and analyzed with an Agilent 6890N chromatograph equipped with an HP-5ms capillary column (50 m by 0.25 mm) and a mass-selective detector [60]. The gas chromatogram conditions were as follows: the injector temperature was set at 280C, the transfer line temperature was 290C, and the flow rate was 1 ml/min with helium. The temperature program of the oven was 104C for two minutes, and then increased to 40  290C at a rate of 15C per minute, then held at 290C for 15 minutes. The mass spec conditions were set to electron emission scanning at 40 – 800 m/z, at 1.97 scans per second.  41  Chapter 3: Results  3.1  RHA1 Cholic Acid Growth Phenotype  RHA1 was grown in M9 mineral medium with 2.0 mM cholic acid as the sole carbon and energy source as described in section 2.2. After a lag phase of roughly 24 hours, RHA1 achieved a specific growth rate of 0.105 h-1 (a doubling time of 2.867 h). At approximately 52 hours, RHA1 reached a maximal OD600 of 1.76  0.02 (Figure 3.1) and a final growth yield of 17.4  0.3 g of protein per 1 ml of culture (Figure 3.2).  2 1.8 Optical Density 600 nm  1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0 0  20  40  60  80  100  120  140  Hours  Figure 3.1 Growth of RHA1 on 2.0 mM cholic acid as the sole carbon and energy source; n=3; error bars show standard deviation.  42  20 18  Final Protein - ug/ml  16 14 12 10 8 6 4 2 0 Wild Type  ∆camB  ∆camBc  ∆camM  ∆camMc  Control  Strain  Figure 3.2 Final protein yield of all strains grown on 2.0 mM cholic acid, control was without inoculum; n=3; error bars indicate standard deviation.  RHA1 was grown on 0.2 mM, 2.0 mM, and 5.0 mM cholic acid, in M9 mineral medium, to assess the effect of cholic acid concentration on growth kinetics. With 0.2 mM, compared to 2.0 mM, cholic acid, RHA1 had a shortened lag phase of 15 hours but grew to a lower final OD600 of 0.30  0.02. The 5.0 mM cholic acid cultures failed to grow (data not shown), presumably due to toxicity of cholic acid. Thus, it was determined that 2.0 mM is the optimal experimental concentration for RHA1 to achieve maximal growth on cholic acid.  43  3.1.1  Extracellular Cholic Acid Metabolite Accumulation by RHA1  Three main cholic acid metabolites were detected using GCMS analysis: 12-hydroxy9-oxo-1,2,3,4,10,19,23,24-octanorcholan-5,22-dioic acid (HOONCA) (Figure 3.3), 12hydroxy-9-oxo-1,2,3,4,10,19,23,24-octanorchol-6-en-5,22-dioic acid (6-HOONCA) (Figure 3.4), and 3,7,12-trihydroxy-9-oxo-9,10-seco-23,24-bisnorchola-1,3,5(10)-trien-22-oic acid (THOSBNC) (Figure 3.5). At roughly 36 hours after inoculation, these compounds reached their maximum level of accumulation in the supernatant, corresponding an estimated 12.5%, 4%, and 3%, respectively, of the initial cholic acid added ( 0.816 g/L), based on the gas chromatogram peak areas.  Figure 3.3 Cholic acid metabolite HOONCA.  Figure 3.4 Cholic acid metabolite 6-HOONCA.  44  Figure 3.5 Cholic acid metabolite THOSBNC.  By 45 hours post-inoculation, less than half of the above compounds were still present in the supernatant of wild type RHA1, and by late stationary phase these compounds were completely reassimilated and are no longer present in the supernatant. Other metabolites were also found to transiently accumulate in the supernatant, yet they were in such low abundances compared to the above compounds, that they were only considered minor metabolites.  3.2  Expression of the ATP-Binding Cassette and MFS Transporters  In a previous study, transcriptomic analysis of RHA1 was performed in which many genes were found to be upregulated during exponential growth on cholic acid, compared to growth on pyruvate (W.W. Mohn, unpublished). Among these upregulated genes were genes encoding two transporters: a four-gene operon encoding an ATP-binding cassette (ABC) transporter, CamABCD (ro04888 – ro04885), and a major facilitator superfamily (MFS) transporter, CamM (ro05792). The prefix, Cam, signifies Cholic acid metabolite transporter. The transcriptomic data revealed that during mid-log phase, the gene encoding a permease component of the ABC-transporter (camB) exhibited 7.4-fold upregulation, and the MFS 45  transporter gene (camM) showed 26.6-fold upregulation, when RHA1 was grown on cholic acid compared to pyruvate. To verify upregulation indicated by the transcriptomic data, RT-QPCR was performed on camB as well as camM. The camB gene was most highly upregulated during early log phase, with a normalized 16.7-fold change in transcription, then still upregulated but to a lesser extent in late log phase with an 8.9-fold change in transcription. By early stationary phase, RHA1 grown on cholic acid exhibited almost no difference in levels of transcription of camB compared to RHA1 grown on pyruvate (Figure 3.6).  18 Fold change compared to pyruvate  16 14 12 10 8 6 4 2 0 early log  late log  early stationary  Stage of Growth  Figure 3.6 Relative upregulation of camB during growth on cholic acid based on RT-QPCR.  46  Figure 3.7 The ABC-transporter operon genes: camA, substrate binding domain; camB, permease component; camC, fused permease/ATP-binding domain; camD, ATP-binding domain, the arrow indications the direction of transcription.  200  Fold change compared to pyruvate  180 160 140 120 100 80 60 40 20 0 early log  late log  early stationary  Stage of Growth  Figure 3.8 Relative upregulation of camM during growth on cholic acid based on RT-QPCR.  Levels of camM transcription were 38.83-fold and 173.7-fold higher during early log and late log phase, respectively, for RHA1 grown on cholic acid compared to pyruvate. Again, during early stationary phase, there was almost no difference in transcription levels of camM between RHA1 grown on cholic acid and pyruvate (Figure 3.8). These RT-QPCR data  47  confirm that both transport systems are significantly upregulated when RHA1 is grown on cholic acid, suggesting that they are important for cholic acid catabolism.  3.3  Inactivation of the ABC Transporter  CamABCD was inactivated by in-frame deletion of camB as described in section 2.4 (Figure 3.9). The permease gene was selected for deletion, as it is the best way to ensure that a single gene deletion inactivates the ABC transporter. The substrate-binding domain and ATP-binding domains of ABC transporters have been found to be somewhat “promiscuous” and can function with other ABC transport systems [61].  Figure 3.9 Colony PCR electrophoresis gel screening for camB transconjugation: the clones represented in lanes A and D have a truncated copy of camB (480 base pairs), lanes B, C and E have a wild type copy (1000 base pairs), the lane D clone was selected for subsequent growth experiments.  48  3.3.1  The Cholic Acid Growth Phenotype of camB  The camB mutant was grown on 2.0 mM cholic acid in the same conditions as the wild type. Like wild-type RHA1, camB exhibited an initial lag phase of 24 hours, followed by exponential growth in which the strain achieved a specific growth rate of 0.094 h-1 with a doubling time of 3.20 hours. The camB mutant reached a maximum OD600 of 1.47  0.01 at 41 hours after inoculation, and then dropped to 1.40  0.02 and slowly decreased in OD600 until late stationary phase (Figure 3.10). The final protein yield of camB was 12.9  0.02 g/ml, which is 74% of the final protein yield of wild type RHA1 (Figure 3.2). Thus, growth kinetics of the mutant were very similar to the wild-type, but the final cell yield was lower.  2 1.8  Optical Density 600 nm  1.6 1.4 1.2 1 RHA1  0.8  ∆camB  0.6 0.4 0.2  0 0  20  40  60  80  100  120  140  Hours  Figure 3.10 camB cholic acid growth phenotype compared to wild type; n=3; error bars indicate standard deviation.  49  3.3.2  Extracellular Cholic Acid Metabolite Accumulation by camB  Extracellular metabolites were extracted from camB cultures at various times, and it was found that this mutant strain accumulates the same metabolites as wild type. However, unlike wild type RHA1, camB was unable to reassimilate two of these metabolites: HOONCA (Figure 3.3) and 6-HOONCA (Figure 3.4). The maximum accumulation of these HOONCA metabolites roughly corresponds to the point of maximum growth of camB, just after 40 hours (Figure 3.11). These metabolites were identified by their retention time (Figure 3.12), fragmentation pattern, and molecular ion (3.13 and 3.14), as detected by the GCMS, which were consistent over repeated experiments. The structures were determined based on studies of other cholic acid metabolizing microorganisms [45], as well as structures identified in studies of RHA1 grown on cholesterol [23].  50  25  Normalized Peak Abundance  20 15 HOONCA  10  D6-HOONCA 5 0 38.5 Hours -5  45 Hours  111 Hours  Time Since Inoculation  A. 30  Normalized Peak Abundance  25 20 15 HOONCA 10  D6-HOONCA  5 0 38.5 Hours  45 Hours  111 Hours  Time Since Inoculation  B.  Figure 3.11 Accumulation of HOONCA metabolites in (A) wild type RHA1 versus (B) camB, based on GCMS normalized peak abundance; n=3; error bars indicate standard deviation.  51  Abundanc e T IC: 9901002.D 14.60  3.5e+07  3e+07  2.5e+07  2e+07  14.83  1.5e+07  1e+07  5000000  16.26  6.00  8.00  10.00  12.00  14.00  16.00  18.00  20.00  22.00  24.00  26.00  28.00  T ime-->  Figure 3.12 Chromatogram of camB stationary phase supernatant showing peaks of accumulated metabolites. Rt 14.60: HOONCA, rt 14.83: 6-HOONCA, rt 16.26: cholestane internal standard.  Figure 3.13 Mass spectrum of HOONCA.  52  Figure 3.14 Mass spectrum of 6-HOONCA.  Figure 3.15 Color-comparison of supernatants: A. camB, B. Wild Type, C. camM.  The supernatant of camB also exhibited a noticeable difference in color when compared to wild type (Figure 3.15).  53  3.3.3  RHA1 and camB Growth on HOONCA Metabolites  0.4  Optical Density 600 nm  0.35 0.3 0.25 0.2  Wild type ∆camB  0.15  uninoculated control  0.1 0.05 0 0  5  10  15  Days  Figure 3.16 Growth of RHA1 and camB on HOONCA metabolites as a carbon and energy source; n=3; error bars indicate standard deviation.  RHA1 and camB were pre-cultured on 2.0 mM cholic acid, then cultured on HOONCA plus 6-HOONCA, harvested from camB supernatant, as the major sources of carbon and energy (Figure 3.16). After seven days, wild type RHA1 entered exponential growth, reaching a maximum OD600 of 0.35  .02 at around day 13. However camB exhibited no growth on these metabolites, reaffirming that the inactivation of this ABCtransporter inhibits the assimilation and catabolism of these HOONCA compounds.  54  3.4  Inactivation of the MFS Transporter  CamM was inactivated by in-frame deletion of the camM gene as described in section 2.4 (Figure 3.17).  Figure 3.17 Colony PCR electrophoresis gel screening for camM transconjugation: the clone represented in lane A has a wild type copy of camM (1700 base pairs), lane B has a truncated copy (900 base pairs) and was used in subsequent growth experiments.  3.4.1  The Cholic Acid Growth Phenotype of camM  On cholic acid, camM exhibited a specific growth rate of 0.074 h-1 with a doubling time of 4.068 hours during exponential growth. The camM mutant appeared to reach the same maximum OD600 as wild type RHA1 (Figure 3.18), however this was not a faithful representation of total biomass produced. The camM mutant only reached a final growth yield of 13.6  0.03 g/ml protein, which is 77.2% the final protein yield of wild type.  55  2  Optical Density 600 nm  1.5  1 Wild Type Average ∆camM  0.5  0  0  20  40  -0.5  60  80  100  120  Hours  Figure 3.18 Growth of camM on 2.0 mM cholic acid, compared to wild type, measured via OD 600; n=3; error bars indicate standard deviation.  The discrepancy between optical density and growth yield is likely due to absorbance at 600 nm of compounds in the supernatant which resulted in the visible dark color (Figure 3.15). The supernatant of camM without cells has an OD600 of 0.085, while the cell-free supernatant of RHA1 has an OD600 of 0.021. This absorbance may be due to a transformation product of the metabolite or to other material released by camM. To get an accurate growth curve for camM, a protein assay was performed to calculate the growth yield at selected points in the growth curve and compare it to that of wild type (Figure 3.19). The protein assay confirmed that camM does not exhibit the same growth kinetics as wild type, despite the similar OD600 growth curves.  56  20  ug protein/mL  15  10 RHA1  ∆camB 5  ∆camM  0 0 -5  20  40  60  80  100  Hours  Figure 3.19 Growth of wild type RHA1 and the transporter mutants on 2.0 mM cholic acid, using protein as a measurement of biomass; n=3; error bars indicate standard deviation.  3.4.2  Extracellular Cholic Acid Metabolite Accumulation by camM  The camM strain accumulated the same metabolites as wild type. However, camM accumulated THOSBNC (Figure 3.5) to a higher extent than wild type, was unable to reassimilate this compound, and maintained it in the supernatant into late stationary phase (Figure 3.20). This metabolite was identified by its retention time, fragmentation pattern, and molecular ion, as detected by the GCMS, which were consistent over repeated trials (Figure 3.21 and Figure 3.22).  57  3.5  Normalized Peak Abundance  3 2.5 2 RHA1 1.5  ∆camM  1 0.5 0  110 Hours  Figure 3.20 THOSBNC accumulation in RHA1 and camM based on normalized GCMS peak abundance; n=3; error bars indicate standard deviation.  Abundanc e T IC : 0 6 0 1 0 0 6 .D 1 8 .9 5 1e+07 9000000 8000000 7000000 6000000 1 5 .9 8 5000000 4000000 3000000 2000000 1000000  6 .0 0  8 .0 0  1 0 .0 0  1 2 .0 0  1 4 .0 0  1 6 .0 0  1 8 .0 0  2 0 .0 0  2 2 .0 0  2 4 .0 0  2 6 .0 0  2 8 .0 0  T im e -->  Figure 3.21 Chromatogram of camM stationary phase culture showing peaks of accumulated metabolites. Rt 15.98: cholestane internal standard, rt 18.96: THOSBNC, other peaks are minor metabolites with similar fragmentation patterns as THOSBNC.  58  Figure 3.22 Mass spectrum of THOSBNC.  3.4.3  RHA1 and camM Growth on THOSBNC  RHA1 and camM were incubated on THOSBNC, harvested from camM supernatant, as the major source of carbon and energy, as described in section 2.2 (Figure 3.23). Wild-type RHA1 cells entered exponential growth beginning around three days after inoculation, reaching a maximum OD600 of 0.09 at around day 8. However camM exhibited no growth on these metabolites, reaffirming that without the inactivated MFS-transporter, camM is unable to reassimilate and catabolize THOSBNC.  59  0.1 0.09 Optical Density 600 nm  0.08 0.07 0.06 0.05  Wild Type  0.04  ∆camM  0.03  uninoculated control  0.02 0.01 0 0  2  4  6  8  10  12  14  Days  Figure 3.23 Growth of RHA1 and camM on the THOSBNC metabolite as a carbon and energy source; n=3; error bars show standard deviation.  3.5  Cholic Acid Conjugate Growth Phenotype of camM  The camM mutant was grown on the conjugated bile acids glycocholic acid and taurocholic acid (Figure 3.24 A and B respectively) to see if disrupting camM had an effect on the uptake of cholic acid in the conjugated form. On taurocholic acid, camM exhibited the same growth kinetics and final growth yield as observed during growth on cholic acid, and also accumulated THOSBNC without reassimilating the compound. However, growth on glycocholic acid was significantly slower, taking approximately five days instead of two days to reach stationary phase (data not shown). The final growth yield on glycocholic acid was 17.0 g/ml (±0.3), which is approximately equal to the wild type. Finally, there were no accumulated metabolites in the supernatant of stationary phase camM cultures grown on  60  glycocholic acid. The camB mutant showed the same growth kinetics and metabolite accumulation in stationary phase from both conjugated bile acids and cholic acid.  A.  B.  Figure 3.24 Structures of conjugated bile salts: A. glycocholic acid, B. taurocholic acid.  3.6  Complementation  The camB and camM mutants were complemented by insertion of the corresponding genes with their native promoters into the chromosome of the mutant strains. For camB, the full camABCD operon was inserted to avoid any issues with protein complex assembly or polar effects on gene expression. Since camM is not part of an operon, only the single gene was inserted for complementation. Insertion complementation was performed in order to avoid any issues with over-expression. Because camM and camABCD encode membrane proteins, the expression of too many copies may have resulted in unforeseen effects on cellular functioning, which can occur when using a high copy-number expression vector like pTIP. Both the complementation strains, named camBc and camMc, were grown on cholic acid in the same manner as the other strains in this study. Both strains grew slower  61  than the wild-type on cholic acid. A final protein assay showed camBc and camMc reached 98.5% and 89.0%, respectively, of the final growth yield of wild type RHA1 (Figure 3.2). Thus, for both mutants, complementation substantially restored the wild-type cell yield on cholic acid. The somewhat lower yield of each complementation strain may be related to the slower growth rate. Both strains produced the same metabolites as wild type RHA1 and also, like the wild type, reassimilated these metabolites such that no compounds were detected in the supernatant of stationary phase cultures. Thus, cloning in a wild-type copy of the respective inactivated transporter gene for both camB and camM was sufficient to restore each strains’ capacity to uptake all extracellular metabolites. This confirms that CamABCD is required for uptake of the HOONCA metabolites, and CamM is required for uptake of THOSBNC.  62  Chapter 4: Discussion  4.1  Cholic Acid Uptake and the Potential for Passive Diffusion  Originally, I hypothesized that one of the two upregulated transporters could be required for the uptake of cholic acid across the RHA1 cytoplasmic membrane. In order to be competitive in a natural environment, soil bacteria like RHA1 generally need transporters that give them high affinity for substrates. Specificity for these substrates may vary from narrow (i.e. a single compound) to broad (classes of compounds). The Mce4 system, required for cholesterol but not cholic acid uptake [33], supports the hypothesis of a specific cholic acid uptake system, yet we failed to identify one in this study. As no other transporters were highly upregulated on cholic acid, uptake may occur by a constitutively expressed transporter, perhaps one with broad substrate specificity. Alternatively, it is possible that cholic acid enters RHA1 via passive diffusion across the cytoplasmic membrane. There is some evidence that cholic acid enters bacteria by this mechanism. The alpha-orientation of all the hydroxyl groups on cholic acid creates distinct hydrophobic and hydrophillic parts of the molecule, which allows it to behave as a detergent [4]. One study found a correlation between hydrophobicity and the rate at which bile salts “flip” across egg phosphatidylcholine vesicles, suggesting that deoxycholic acid would traverse the membrane faster than cholic acid since it is more hydrophobic. This study also concludes that all bile salts would flip through the membrane eventually [62]. However, they also predict that cholesterol would flip faster than any of the bile salts, which seems  63  inconsistent with the requirement for the Mce4 transporter for growth on cholesterol [33]. Thus it is unclear whether membrane flipping occurs during bile salt catabolism in RHA1. The only described bacterial bile salt uptake transporter is from the intestinal bacterium Clostridium scindens, called BaiG. This protein catalyzes the proton-motive-forcedependent uptake of bile salts [43]. A putative bile salt uptake transporter gene has been found in Lactococcus johnsonii [63] but neither this gene nor BaiG have significant similarities to putative transporters in C. testosteroni or RHA1 [4]. Thus, it remains unclear whether RHA1 or related organisms employ a transporter for cholic acid uptake, or whether it merely diffuses into the cell.  4.2  A Unifying Scheme for Steroid Degradation  Transient extracellular metabolite accumulation during cholic acid degradation is a general phenomenon seen in diverse microorganisms. I found that RHA1 is no exception. The proposed structures of the RHA1 cholic acid metabolites are similar to those detected in other organisms [4, 24, 45]. What is novel about the RHA1 metabolites is that they retain an isopropionyl group at C17, whereas other metabolites detected in previous studies have a ketone group at this position. This suggests that side-chain degradation may reach completion later in the overall degradation pathway in RHA1 than it does in other cholic acid degrading bacteria. Despite this distinction, the RHA1 metabolites identified here are consistent with the common pathway of microbial cholic acid degradation [22]. The formation of the secosteroid THOSBNC suggests that the degradation of rings A and B happens in a similar  64  fashion in RHA1 as it does in organisms like strains TA441 and Chol1 [24, 45]. In addition, the formation of the HOONCA metabolites also agrees with the seco-steroid pathway constructed from the degradation of other steroid compounds, which all have DOHNAA as the final compound identified before the unknown final steps leading to CO2 [24]. These metabolites further suggest that the hydroxyl group at C7 is not metabolized until after cleavage of Ring B, and that at C12 is not metabolized until after removal of Ring A. Results here support the hypothesis that despite different side chain conformations and the attachment of hydroxyl groups at various locations on the steroid nucleus, the degradation of most steroids follows the aforementioned seco-steroid pathway. The main differences appear to lie in the order of side chain degradation relative to A-ring oxidation and B-ring opening, which varies among organisms studied and the steroids degraded. This study was the first to demonstrate the function of putative cholic acid genes through targeted mutagenesis. Compared to cholesterol and testosterone, the cholic acid pathway is not as well-understood. No catabolic enzymes have yet been purified or characterized. The requirement for transporters involved in cholic acid metabolite uptake has been suggested in previous publications [45], but this study provides the first evidence for and identification of such transporters.  4.3  Support for the Toxic Metabolite Excretion Hypothesis  Like other microorganisms in previous studies, I’ve demonstrated that RHA1 also transiently accumulates extracellular metabolites during growth on cholic acid. A hypothesis frequently stated is that these metabolites represent precursors to rate-limiting steps in the  65  cholic acid pathway. At this “bottle neck” in the catabolic pathway, the excess metabolites are excreted into the extracellular environment until the cell is ready to incorporate them back into the cell and further degrade them. This process, including the transporters used in assimilation of the metabolites, would be an adaptive strategy employed by diverse bacteria to maintain low intracellular levels of these metabolites, which are potentially toxic to the cell [45], while maximizing nutritional benefit from cholic acid. I found additional support for this hypothesis in the growth phenotype on glycocholic acid of camM, which took longer to reach stationary phase than the wild type, yet reached the same final growth yield and did not accumulate THOSBNC. I hypothesize that the initial steps in degradation of glycocholic acid are slower than for cholic acid, due to the altered conformation of the side chain, resulting in a reduced accumulation of THOSBNC, thus eliminating the need for its excretion. This lack of metabolite accumulation was only observed during growth on glycocholic acid of camM, and not of camB, which showed the same metabolite accumulation in stationary phase from both conjugated bile acids and cholic acid. This could be because camB is required to transport metabolites formed later in the pathway, at which point the rate of metabolite generation may be equal for all bile acid substrates. Finally, efflux transporters for the excretion of these metabolites likely exist. It is possible that these transporters have broad specificity and have a general function in removal of potentially toxic compounds from the bacterial cell. Such transporters would likely be constitutively expressed.  66  4.4  Two Novel Transporters  Both the ABC and MFS superfamilies are highly diverse; they include transport systems that operate in a range of organisms, from bacteria through humans, to pump a wide variety of substances across biological membranes. Substrate specificity varies greatly, from very specific to very broad, depending on the transporter and the substrate(s) involved [64, 65]. Despite the progress of research on ABC and MFS transporters, many gaps exist in the understanding of their structure and function [64, 66]. Genome sequences have revealed vast numbers of these transporters with unknown functions. There is great need to assign function to transporters in both families in order to advance our general understanding of these important micro-machines. This study contributes to the general knowledge of both the ABC and MFS superfamilies by identifying novel members of each superfamily with defined transport functions. Further studies on the structures of the CamABCD and CamM systems, as well as investigations on the specificity of each, will contribute to the elucidation of these unique transporters. In particular, since these transporters have novel substrates, they may contribute to understanding the relationship between protein structure and substrate specificity.  4.5  Habitat of Cholic Acid Degraders  This study may provide insight into the environments to which RHA1 has adapted during its evolutionary history. One might assume that RHA1, as well as other free-living microorganisms that can catabolize cholic acid, would have adapted to sequester this  67  compound from environments in which cholic acid is present in very minute amounts. However, the strategy of metabolite efflux observed here suggests that RHA1 has evolved to deal with cholic acid in much higher concentrations than the limited amount that would likely be present in a given soil sample. Only in environments with a high cholic acid concentration, like the cultures used for this study, would such an efflux process be necessary. This suggests that feces may be a significant habitat for RHA1 and other cholic acid-degrading organisms.  4.6  Implications  Better understanding of steroid metabolite accumulation and uptake may have applications in an industrial context, as this phenomenon may affect the use of whole cell biocatalysis in the manufacture of steroid-related therapeutics. Production of toxic metabolites or metabolite reassimilation could potentially be rate-limiting for a manufacturing process. On the other hand, excretion of desired products could increase the efficiency of a process. Additionally, there may be environmental implications, as the excretion of metabolites might allow receiving environments to rapidly degrade large quantities of certain steroids discharged by municipal or industrial sewage. The metabolites may have environmental effects and should be studied in this context. Finally, further studies on the enzymology involved in cholic acid catabolism in RHA1 are required to fully understand the overall catabolic pathway. These accumulated  68  metabolites could be important and easily available substrates for future activity assays, as they are relatively pure in the supernatant of the camB and camM mutants.  69  References  1.  Hogg JA: Steroids, the Steroid Community, and Upjohn in Perspective - a Profile of Innovation. Steroids 1992, 57(12):593-616.  2.  Ying GG, Kookana RS, Ru YJ: Occurrence and fate of hormone steroids in the environment. Environ Int 2002, 28(6):545-551.  3.  Streck G: Chemical and biological analysis of estrogenic, progestagenic and androgenic steroids in the environment. Trac-Trend Anal Chem 2009, 28(6):635652.  4.  Philipp B: Bacterial degradation of bile salts. Appl Microbiol Biot 2011, 89(4):903915.  5.  Johnston JO: Aromatase inhibitors. Crit Rev Biochem Mol 1998, 33(5):375-405.  6.  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