I A 5:. < v. “fly-w— . Int 1.. UTA» pl. : . . wank.) 50.13.33.125. .....t.v I1! \Jazal 3 ' '[ y'gffff-{g’ I 6. AA .15., to .v ()0 o vv. ILA.‘ . phi..li...a.l . .. . L l l .\ Jl‘) 3 51/;an 34 LIBRARY Michigan State University This is to certify that the thesis entitled CHARACTERIZATION OF THE 188 ss-rDNA OF RHINOSPORIDIUM SEEBERI RESOLVES A CENTURY OF TAXONOMIC UNCERTAINTY. presented by Roger Alan Herr has been accepted towards fulfillment of the requirements for the MS. degre in Clinical Laboratory Sciences 14% Major Professor's Signature 07 /3/ /0 3 Date MSU is an Affinnative Action/Equal Opportunity Institution PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE 6/01 c-JCIRC/Datoouobss-sz CHARACTERIZATION OF THE 188 ss-rDNA OF RHINOSPORIDIUM SEEBERI RESOLVES A CENTURY OF TAXONOMIC UNCERTAINTY. By Roger Alan Herr A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTERS OF SCIENCE Clinical Laboratory Sciences Program 2003 ABSTRACT CHARACTERIZATION OF THE 188 ss-rDNA OF RHINOSPORIDIUM SEEBERI RESOLVES A CENTURY OF TAXONOMIC UNCERTAINTY. By Roger Alan Herr For the past 100 years Rhinosporidium seeben‘ has been classified either as a protozoan or as a member of the Kingdom Fungi. We have amplified and sequenced 1791 base pairs of R. seeben‘ 18$ ss-rDNA. Using phylogenetic analysis, by parsimony and distance methods of the 188 ss-rDNA of R. seeberi and other eukaryotes, we found that this enigmatic human and animal pathogen clusters with a novel group of fish parasites referred to as the “DRIP” clade, near the animal-fungal divergence. Our phylogenetic analyses also indicate that R. seeben’ is the sister taxon of the two Dermocystidium species used in this study. This molecular affinity is remarkable because members of the genus Dermocystidium also form spherical structures in infected hosts, produce endospores, are intractable to culture, and possess flat mitochondrial cn‘stae. With the addition of R. seeberi to this clade, the acronym “DRIP” is no longer appropriate. We propose to name this monophyletic clade the Mesomycetozoa to reflect the group’s phylogenetic association within the Eukarya. DEDICATION I would like to dedicate this body of work to all of the people in my life who directly or indirectly supported me during my time at Michigan State University. Special recognition needs to be given to Wilbur and Jonnie Herr, Douglas Estry, John Gerlach, and Leonel Mendoza for their unwavering belief in my ability to succeed as a student and scientist. ACKNOWLEDGMENTS I would like to acknowledge the Medical Technology Program faculty and staff for their support. TABLE OF CONTENTS DEDICATION - - _ . III ACKNOWLEDGMENTS ...... ‘ - ‘IV TABLE OF CONTENTS V Wis. ~ V11 LIST OF FIGURES - _ - - _ VIII INTRODUCTION - - _ 1 w -- 3 LITERATURE REVIEW ................................................................................................ 3 History of Rhinosporidium seeberi. ............................................................................ 3 The life cycle of Rhinosporidium seeberi within host tissue. ...................................... 4 Nomenclature of Rhinosporidium seeberi ’s in vivo developmental stages ................. 4 Culture. ....................................................................................................................... 8 Pathogenicity in experimental animals ....................................................................... 8 Ecology. ...................................................................................................................... 9 Distribution of rhinosporidiosis .................................................................................. 9 Clinical features ofrhinosporidiosis. ............................ 12 Diagnosis and treatment of rhinosporidiosis ............................................................ 13 Studying Rhinosporidium seeberi at the molecular level. ........................................ 14 The 18S rRNA molecule. ........................................................................................... I 5 CHAPTER 2 -- - - 17 MATERIALS AND METHODS .................................................................................. 17 Identification oLRhinosporidium seeberi by microscopy. ........................................ 1 7 Vectors and Markers. ................................................................................................ 1 7 Media. ....................................................................................................................... 1 7 Buflers. ...................................................................................................................... 18 Light and transmission electron microscopy {T EM) materials. ............................... I8 Moleculgfliology materials. .................................................................................... 19 Collection and storage of tissues containing R. seeberi sporangia gnd endospores.21 Transmission electron microscg sggdies on rhinosporidial tissues. ..................... 22 Molecular studies on rhinosporidifial tissues. ............................................................ 23 CHAPTER 3 - - 38 RESULTS ..................................................................................................................... 38 CHAPTER 4 _ 61 DISCUSSION ............................................................................................................... 61 APPENDIX A A 71 PHYLOGENETIC ANALYSIS OF RHINOSPORIDI UM SEEBERI ’S 188 SSU RDNA GROUPED THIS PATHOGEN AMONG MEMBERS OF THE LROTOCTISTAN MESOMYCETOZOA I‘DRIP’I CLADE ................................. 71 APPENDIX B 84 LOCALIZ_ATION OF AN IMMUNOREACTIVE INNERWALL ANTIGEN OF RHINOSPORIDI UM SEEBERI EXPRESSED ONLY DURING MATURE SPORANGIAL DEVELOPMENT ............................................................................ 84 APPENDIX C 96 IN VITRO STUDIES ON THE MECHANISMS OF ENDOSPORE RELEASE IN RHINOSPORIDI UM SEEBERI ................................................................................. 96 BIBLIOGRAPHY - - 1 l 0 vi LIST OF TABLES TABLE 1: NOMENCLATUE OF R. SEEBERI’S IN VIVO DEVELOPMENTAL. STAGES PROPOSED BY K_ENNEDY ET AL. (ADAPTED FROMLQENNEDYiBUGGAGE AND AJELLO 1995 I. ............................................................................................................ 7 TABLE 2: ABBREVIATIONS AND SEQUENCES OF 18S SSRmA FORWARD PRIM_ERS. ..... 28 TABLE 3: ABBREVIATIONS AND SEQUENCES OF 18S SSRDNA REVERSE PRIMERS. ...... 28 TABLE 4: A_BI PRISMTM BIG DYETM TERMINATOR CYCLE S_E<__2UENCING REACTION. ....... 35 vii LIST OF FIGURES FIGUREr1: QEOGRAPHIC le§UTION OF RHINOSPORIDIOSIS. .................................. 11 FIGURE {MRIMER MAP OF THE 18§R§NA MOLECULE WITH THE LOCATIONS OF NS_1,_ 2. 3. 4. 5. 6. 7. AND N88 DEGENERATE PRIM_ERS (GARGAS ANQQEPREIST 1996). ..................................................................................................................... 29 FIGURE 3: AN 0.8% AGAROSE GEL SHOWING A 1:10 DILUTION AND UNDILUTED NUCLEIC ACIDS OF R. SEEBERI PRIOR To R_NASE TREATMENTS IN LANES 2 AND 3 RESPECTIVELY. .............................................................................................. 40 FIGUR_E 4: THIS FIGURE SHOWS A 0.8% AGAROSE GEL SHOWING R. SEEBERI DNA AFTER THREE RNASE DIGESTS. LANES 2. 3 AND 4 CONTAIN 5,11. OF SAMPLE DNA FROM SEPARATE RNASE TREATMENTS. ............................................................ 41 FIGURE 52 THIS FIGURE SHOWS A 1.2% AGAROSE GEL USED TO DOCUMENT THE PRESENCE OF THE AMPLICON OF PREDICTED S|_ZE PRIOR TO CLONING .................. 44 FIGURE 6: CRACKING GEL #1. (CLONES 1-15) WAS RUN FOR 3 HOURS AT 110V.J=ANE #1 CONTAINS THE NEGATIVE CONTROL (P0321 VECTOR WITH No INSERT). ........ 47 FIGURE 7: CRACKING GEL #gICLONES 16-30) WAS RUN FOR 3 HOURS AT 110V. LANE 16 IS THE NEGATIVE CONTROL (PCRg.1 VECTOR WITH NO lNSER'Il ................... 48 FIGflE 8: CRACKING GEL #3 (CLONES 30-44) WAS RUN FOR 2 HOURS AT toowNE 33 CONTAINS A NEGATIVE CONTROL (PCRQ VECTOR WITH No INSERT). LANE 30 CONTAINS CLONE #30 WHICH WAS REPEATED To CONFIRM THE RESULTS OF CRACKING GEL #2 CLONE 30. ......................................................................... 49 FIGURE 9£MCKING G; #4 (CLONES 45-49 AND 54-63) WA§ RUN FOR 2 HOURS AT 100V. HNES 45 AND 61 CONTAIN N_EGATIVE cgNTROLS (PCR 2.1 VECTOR WHICH @NTAINS NO INSERT). .................................................................................... 50 FIGUR_E 10: THE FIGURE DEPICTs A 529 BASE PAIR FRAGMENT (BASES 1196-1 724) OF THE COMPLETE 1.791 AMPLICON FROM R. SEEBERI’S 18S RDNA MOLECULE. ..... 53 FIGUR_E 11: RHINOSPORIDIUM SEEBERI’S 188 SSRDNA (1791 BASE PAIRS). ............. 54 FIGURE 12: PHYLOGENETIC ANALYSIS BY NEIGHJBOR JOINING ANg PARSIMgNY. WITH R. SEEBERI AND 23 OTHER 18S SSU RDNA SEOUENCES. SUPPORTED VERY SIMILAR PHYLOGENETIC TREES. ................................................................................... 59 viii FIGUR_E 13: THE DIAGRAMS ABOVE ILLUSTRATE ALTERNATIVE RELATIONSHIPS AMONG THE DRIP’S CLADE AND OTHER EUKARYOTIC LINEAGES NOT REJECTABLE UNDER THE TEMPLETON-FELSENSTEIN AND K_ISHINO-HUSEGAWA TESTS ........................ 65 INTRODUCTION Rhinospon'dium seeberi, the etiological agent of rhinosporidiosis, was first observed by Malbran and Seeber(see Ashworth 1923). More than a century has passed and it’s taxonomic position has not yet been resolved. There are several reasons why R. seeberi’s taxonomic status has been difficult to establish: i) it is intractable to culture, ii) it can be found only in naturally infected hosts, iii) nothing is known about its life cycle outside of the host, iv) the geographic distribution of rhinosporidiosis makes tissue samples for molecular studies very difficult to obtain. For these reasons, early studies of R. seeberi were limited to light and electron microscopic examination of infected tissue (Ashworth 1923, Bader and Grueber 1970, Vanbreuseghem 1973, Kannan-Kutty and Teh 1974, and Savino and Margo1983). As a result, the Characteristic morphological stages of R. seeberi’s complex life cycle within the hosts’ tissues were well established. Based on its morphological features R. seeberi has been classified as a fungal or a protozoan organism. Rhinosporidium seeberi has also been erroneously identified as a cyanobacterium, and as carbohydrate waste derived from high levels of tapioca ingestion (Ahluwalia 1992). The clarification of R. seeberi’s taxonomy is of significance because it will help to establish its position within the Eukarya. its epidemiology, and its pathogenesis. In an effort to determine its phylogenetic position within the eukarya, this study uses molecular tools to characterize the 188 small subunit ribosomal DNA (188 ss-rDNA) of R. seeberi and to compare its nucleotide sequence with other eukaryotic 188 SS- rDNA sequences. This study will help to answer questions about R.seeberi’s intractability to culture, its possible life cycle in nature, its mode of transmission, as well as, its taxonomic relationship with other members of the eukarya. Chapter 1 LITERATURE REVIEW History of Rhinosporidium seeberi. ‘There is a certain air of romance attached to the name “Rhinosporidium”, arising from its strikingly beautiful developmental stages and its enigmatic nature’ (Thomas, Gopinath and Betts 1956). Rhinosporidium seeberi was first observed by Malbran in a nasal polyp in 1892. He regarded R. seeberi as a sporozoan, but did not publish his findings. In 1900, Seeber examined a nasal polyp and provided the first full description of the organism in his medical thesis (Seeber 1900). Seeber believed that the organism was a sporozoan allied to the Polysporea of the Coccidia, but he did not name the organism. During the same year, Wemicke published more details on the organism described by Seeber and named it Rhinosporidium seeberi. In 1923, Ashworth, in his classical monograph on rhinosporidiosis, concluded that "the nearest relatives of Rhinosporidium are not the Sporozoa but the lower fungi (Phycomycetes) such as the Chytridineae in which, sub-order, near the Olpidiaceae, Rhinosporidium was provisionally placed”(Ashworth 1923). In 1935, Dodge interpreted the sporangia of R. seeben‘ to be multispored asci and classified it as an ascomycetous fungus (Dodge 1935). Carini, in 1940 observed cysts with Spores in skin nodules in frogs and created a new genus Dermospon‘dium. Carini (1940) believed that this newly described organism was closely related to but different from R. seeben‘. The life cycle of Rhinosporidium seeberi within host tissue. It has been proposed that R. seeberi’s life cycle within host tissue is initiated by trauma after exposure to aquatic environments in which R. seeberi is present. Once the organism has implanted in the host tissue it begins to differentiate into a spherical body (sporangium) with a well-defined cell wall. The sporangium enlarges dramatically and thousands of small oval bodies (immature endospores) begin to form. The immature endospores take on a spherical Shape, increase in size, and become mature endospores. Meanwhile, more immature endospores are being formed at the gerrninative zone of the sporangium (Bader and Grueber 1970). When the sporangium matures, a pore forms in its cell wall proximal to the mature endospores. At this stage there is a visible rearrangement and organization of endospores in preparation for release. Mature endospores migrate towards the pore, while immature endospores remain distal to the pore. The sporangium releases its endospores through the pore into the host tissue. The endospores that have been released increase in size and form juvenile sporangia, which enlarge even further and form their own endospores, and the in vivo life cycle is reinitiated. Nomenclature of Rhinosporidium seeberi’s in vivo developmental stages. “In parallel with its disputed taxonomy, a variety of inconsistent and overlapping terms have been applied to the morphological structures of R. seeberi in its developmental stages (Arseculeratne and Ajello 1997)”. Kennedy, Buggage and Ajello (1995) proposed the terminology presented in table 1 which best describes the in vivo developmental stages of R. seeberi. The following are the current nomenclature and definitions used to describe R. seebefi’s phenotypes in infected tissue. Endosmres. Endospores are the asexual form of R. seeberi. A mature sporangium may contain up to 20,000 endospores (Ashworth 1923). Two types of endospores have been observed: immature endospores and mature endospores. 1) Immature endospores range from 2-4 pm in diameter and are formed by cytoplasmic cleavage inside the mature sporangium. They are usually oval in shape and restricted to the portion of the mature sporangium distal to the pore. This region has been referred to as the gerrninative zone of the sporangium (Bader and Grueber 1970). 2) Mature endospores range from 6-10 pm in diameter. These endospores have a well-defined cell wall with a prominent capsule and often have a visible nucleus and several lipid bodies. Mature endospores are only found in mature sporangia that have reached the final stages of development. Juvenile smrangium Mature endospores increase in size and become juvenile sporangia which range in size from 10-100 pm in diameter. The cell wall of a juvenile sporangium is a single electron dense layer. The remnants of capsules from previous stages are usually present on the outer layer of the cell wall. Juvenile sporangia contain a single nucleus, as well as lipid bodies. Cytoplasmic cleavage (endospore formation) has not yet begun at this stage. Intermediate sporangjgm Intermediate sporangia range in size from 100-150 pm in diameter. In this stage the cell wall has thickened. The remnants of a capsule may still be observed. The cytoplasm of an intermediate sporangium is well differentiated, and is characterized by the presence of numerous membrane bound vesicles, mitochondria, nuclei, lipid bodies, and laminar bodies. The cytoplasmic activity during this stage of sporangial development is distinct. All the cytoplasmic changes at this sporangial stage indicate that endospore formation has begun. Mature sporangium Mature sporangium range in size from 150-500 um in diameter. This stage is characterized by the development of one or more inner, mucoid-like, cytoplasmic electron-lucent layers (CEL) located immediately below R. seeberi’s sporangium thin cell wall (Herr et al. 1999). These inner layers are only found in mature sporangium. The mature sporangium contains either immature or mature endospores. A pore develops in the mature sporangium’s cell wall just prior to endospore release. Endospores are released through this pore when the internal pressure of the sporangium increases. Table 1: Nomenclature of R. seeberi’s in vivo developmental stages proposed by Kennedy et aI. (adapted from Kennedy, Buggage and Ajello 1995; Earlier terminology Current terminology Spore, immature spore, endospore, sporoblast, early trophic stage, spherule, conidium, sporule. Mature spore, sporont, pansporoblast, merozoite, yeast phase in tissue, spore morula. Trophocyte (early, intermediate, late), trophic stage, trophozoite, pre-cleavage phase, immature sporangium, granular stage, sporocyst, cyst, spherule, sporangium. Endosporulating stage, sporulation phase, trophic sporangium. Mature trophocyte, mature cyst, endosporulating stage, post- cleavage phase, post-cleavage sporangium, adult stage. Immature endospore Mature endospore Juvenile sporangium Intermediate sporangium Mature sporangium Culture. The etiological agent of rhinosporidiosis has yet to be cultured. Although several investigators claimed to have cultured Rhinosporidium seeberi in vitro, these claims of propagation have not been replicated. A wide variety of bacteriological, mycological, and cell culture media, as well as cell line cultures have been used in an attempt to isolate R. seeberi (Grover, Karunaratne, Krishnamoorthy, Levy 1986). Due to the fact that a mature sporangium contains thousands of endospores that are released Upon contact with water; early investigators believed they had cultured the organism when the release of the endospores was observed in broth cultures. Pathogenicity in experimental animals. In addition to its intractability to culture, attempts to experimentally infect animals with R. seeberi propagules have not been successful. Ajwide range of animals including: cats, goats, guinea pigs, horses, mice, monkeys, rabbits, rats, sheep, snails, and frogs have been experimentally inoculated, but sustained or progressive disease was not observed. Attempts to experimentally infect animals have failed, perhaps due to the lack of information about R. seeberi’s life cycle outside of host tissue. The stage of R. seeben' that is able to infect animals and humans is unknown, which may explain unsuccessful attempts to infect animals with endospores and sporangia as inoculum. Ecology. It is not known whether R. seeberi occurs as a free-living organism or if it has an intermediate reservoir before infecting humans or animals. Due to R. seeberi’s epidemiological features it is believed that this organism is associated with fresh water in ponds, lakes, or in soil. Unfortunately, to date the examination of aquatic animals, silt, manure, water, and aquatic plants from areas where the disease has occurred have not revealed any evidence of R. seeberi (Reddy and Lakshminarayanan 1962). Since R. seeberi has never been cultured or observed outside a host, its morphological features outside of the host are unknown. Thus, the determination of its ecological niche has been difficult establish. Distribution of rhinosporidiosis. Rhinosporidiosis is a disease usually associated with tropical and sub- tropical environments. With the exception of Australia, cases of rhinosporidiosis have been reported in humans and animals of all continents. Sri Lanka and India have the highest prevalence of rhinosporidiosis (Figure 1 page 11). Argentina, Brazil, and the USA (Texas) are also regarded as endemic foci (Karunaratne 1964, Kennedy, Buggage and Ajello 1995). There have been only two reported outbreaks of rhinosporidiosis. The first was noted in humans from northern Serbia from 1992-1993. This epidemic affected 17 people who had been bathing in a stagnant body of water while on vacation in the area. The significance of the epidemic is brought into perspective upon noting that the total number of cases reported in Europe previous to this event barely exceeded one dozen (VukoviC et al. 1995). The other outbreak of rhinosporidiosis was reported in captive swans dwelling on a lake in Central Florida City (FL, USA). Over a three year period 41 captive swans developed rhinosporidiosis (Kennedy et al 1995). In addition, recently a number of captive swans have been diagnosed with rhinosporidiosis in the same lake in Central Florida City (personal communication). The occurrence of the disease in swans supports the belief that R. seeberi is linked to an aquatic environment. Sporadic cases of rhinosporidiosis have also been reported in dogs, ducks, and horses in North America (Mosier, et al. 1984, Davidson, et al. 1977, and Smith, et al. 1961). In addition, a Single case of rhinosporidiosis in a river dolphin from Brazil has been reported (personal communication). 10 D Sporadic rhinosporidiosis. % Frequent rhinosporidiosis. D No reported cases Figure 1: Geographic distribution of rhinosporidiosis. 11 Clinical features of rhinosporidiosis. Rhinosporidiosis is a granulomatous disease of humans and animals characterized by the development of painless slow growing polyps associated with the hosts’ mucous membranes. Rhinosporidial polypoidal masses are soft friable and red in color due to an increase in vascularity. The increase in vascularity in the polypoidal tissue leads to frequent bleeding. In patients with nasal polyps, even forceful sneezing or coughing results in the emission of mucous and blood from the surface of the growth. The small white spots, visible to the unaided eye on the surface of the polyp, are the mature sporangium of R. seeberi. In cases of cutaneous rhinosporidiosis older growths may appear verrucous. Rhinosporidiosis is rarely a life threatening disease. However, a few cases of dissemination have been reported where the infection was fatal (Rajam et al. 1955). The majority of human rhinosporidiosis cases are nasal infections (70-75%), eye infections (15%), and infections of other mucosal areas such as the urethra, vagina, or anus (10%) (Karunaratne 1964). There are also few cases of cutaneous infection (Rippon 1982) and disseminated rhinosporidiosis (Agrawal et al. 1959, Chatterjee et al. 1977, Ho and Tay 1986, Mahakrisnan et al. 1981, Rajam et al. 1955). The dissemination of R. seeberi to anatomically unrelated Sites in the body has been attributed to a haematogeneous route. Arseculeratne (Arseculeratne and Ajello1998) described a case where a patient developed lesions on 20 different sites of the body a few years after surgical excision of a polyp. One such lesion was a large 12 rhinosporidial mass which appeared at the site of a previous closed injury on the leg. More, recently a new case of disseminated rhinosporidiosis was reported in Sri Lanka (Angunawela et al. 1999). This case was in a 44 year old male farmer who presented with a large mass on his left thigh and a second large mass on his sternum. Both masses had been growing slowly for over a two year period. The patient’s clinical history showed that he had been bleeding from the nose for two to three years prior to the development of the two large masses. Both masses were surgically excised without cauterization. The diagnosis of rhinosporidiosis was made after the histopathological examination of the excised masses. During the Six months following the surgical excision the patient developed six more nodules on his trunk and limbs. Diagnosis and treatment of rhinosporidiosis. The Clinical features of rhinosporidiosis mimic several skin and mucous membrane diseases from which it needs to be differentiated. The definitive diagnosis of rhinosporidiosis is made by histopathological examination of infected tissues. The in vivo morphological developmental stages of R. seeberi are easily identified in rhinosporidial tissue with the use of haematoxylin and eosin (H&E) stain. Other stains such as: Periodic acid shiff (PAS), mucicarmine, and Gomori’s methenamine silver (GMS) are useful for diagnostic purposes. Currently there are no serological or nucleic acid based tests to diagnose rhinosporidiosis. Treatment of rhinosporidiosis has been limited to surgical removal of polypiodal masses. Surgical removal of polyps without cauterization may result 13 in dissemination of R. seeberi. It is believed that the dissemination occurs via haematogeneous routes or by contamination of the adjacent mucosal surfaces by endospores (Khan Ali Afzal, Khaleque and Juda 1969). Satyanarayana (1960) estimated that the recurrence rate after surgery was about 11% (Satyanarayana 1960). The use of dmg therapy to treat cases of rhinosporidiosis has been ineffective. Quinine hydrochloride (Wright 1922), salts of antimony and bismuth, iodine and pentamidine (Rajam et al. 1955), griseofulvin, amphotericin B, topical steroids (Jimenez, Young and Hough 1984, Ho and Tay 1986), local or systemic (antibacterial) antibiotics and radiotherapy (Satyanarayana 1966) were all useless for the treatment of rhinosporidiosis. The failure of drug therapy in rhinosporidiosis has been attributed to the impenetrability of the sporangial wall (Woodard and Hudson 1984). However, it is possible that R. seeberi’s phylogenetic connection with the protoctista may have something to do with its resistance to antifungal drugs. Studying Rhinosporidium seeberi at the molecular level. The use of molecular approaches to determine the phylogenetic affinities of Rhinosporidium seeberi has never been successful. However, medical mycologists and eukaryotic phylogenists suggested that a molecular approach to studying this pathogen would be of importance in unveiling its phylogenetic affinities. Kwon-Chung in 1994 stated that, “The phylogeny of the agents of lobomycosis and rhinosporidiosis, although they are treated as fungi, remains unknown. Although there is no in vitro culture system for Loboa Ioboi and Rhinosporidium seeberi at present, a molecular approach would allow us to reveal their phylogenetic relationship, and we can hope that such attempts are forthcoming.” (Kwon—Chung 1994). 14 To study R. seeberi at the molecular level it is necessary to choose a molecule that will be useful as a measure of phylogenetic relationships. Currently, the most comprehensive set of molecular data available for eukaryotes is the 188 serNA sequences. In addition, the 18S molecule has regions that are extremely conserved, slightly variable, as well as regions of great variability. The variation of compositional change within the 18S molecule is what makes it suitable for a wide range of phylogenetic comparisons. The reason why scientists had difficulties characterizing R. seeberi at the molecular level was because it is difficult to obtain a suitable sample to isolate R. seeberi’s nucleic acids or proteins. Due to the location of the endemic areas of rhinosporidiosis, it is difficult to obtain a fresh and unfixed tissue sample in a timely fashion. Another difficulty is the isolation of nucleic acids, and proteins from a complex tissue specimen that contains human or animal tissue, and possibly other microbial contaminants (natural flora) as well as R. seeberi. The 18$ rRNA molecule. The 188 rRNA molecule is transcribed from rDNA that codes for a 45S rRNA primary transcript which is post-transcriptionally modified within the nucleolus to yield the SS, 188, and 288 rRNA’s. The 18S, 28S, and SS rRNA sequences allow the molecules to fold upon themselves extensively and associate with specific proteins to yield ribosomes. The rRNA molecules are necessary as structural components of the ribosome upon which translation actually takes place. One reason why the 18S rRNA is useful as a measure of phylogenetic relationships is because it can be used over a wide spectrum of 15 phylogenetic distances, from kingdom to kingdom at one extreme, to distinctions among species within the same genus. This is possible because the substitution rates at different areas of the 188 molecule vary by at least two orders of magnitude (Olsen 1987, Olsen and Woese 1989). 16 Chapter 2 MATERIALS AND METHODS MATERIALS Identification of Rhinosporidium seeberi by microscopy. Clinical material for this study was obtained from two separate cases of rhinosporidiosis in Sri Lanka. The infected tissue was examined at the light microscopic and electron microscopic levels by Sarath N. Arseculeratne (at the clinical site in Sri Lanka) and Libero Ajello (Emroy University) to confirm the presence of R. seeberi. Upon arrival at the laboratory of Dr. Leonel Mendoza at Michigan State University the tissue sample was examined again using both light and electron microscopic techniques to confirm the presence of R. seeberi. Vectors and Markers. Eschericia coli TOP10 competent cells, and the pCR ® 2.1-TOPO vector were purchased from lnvitrogen (Carlsbad, CA) for cloning of PCR amplicons. A high molecular weight marker (1» Hindlll digest 23.1 kb - 564 bp) was purchased from Bio-Rad (Hercules, CA). A 1 kb molecular weight ladder was obtained from Promega (Madison, WI). Media. Saubouraud agar plates (10 g/L tryptone, 40 g/L glucose, 15 g/L Difco agar) were used to isolate or identify possible contaminants from rhinosporidial tissues. Luria-Bertaini kanamycin broth (LBK) (10 g/L of NaCl, 10 g/L tryptone, 5 17 g/L yeast extract the pH was adjusted to 7.0 with 5N NaOH and 50 pg/mL of kanamycin [Sigma St. Louis, MO] was added to the cooled broth) was used to grow and maintain the pCR® 2.1-TOPO vector in the TOP10 cells. Luria- Bertaini kanamycin (LBK) agar plates were made as the LBK broth above and 20 glL of Difco agar was added to the medium. After the plates were cooled, 40 pL of a 40 pg/mL solution of X-gal (Promega Madison, WI) was spread evenly onto each 100 mm diameter plate. The LBK agar plates were used for putative positive clone selection and maintenance of selected clones. Buffers. The buffers used throughout this investigation were prepared by the standard methods described in (Maniatis et al.). TAE (10X) was prepared as 48.4 g/L tris base, 11.4 mL/L glacial acetic acid, and 20 mL/L of 0.5 M EDTA. LySis buffer was prepared fresh in 20 mL batches as: 2 mL of 50mM Tris-HCL (pH 7.2), 2 mL of 50mM EDTA, 6 mL of 3% SDS, 200 pL of 1% 2- mercaptoethanol, and 9.8 mL of deionized H20. Cracking buffer was prepared fresh in 100 mL batches as: 0.8 g NaOH, 0.5 9 SDS, 20 9 sucrose and enough deionized water to bring the volume up to 100 mL. Light and transmission electron microscopy (TEM) materials. Primary fixation buffer (2.5% glutaraldehyde in 0.05M Na-Cacadylate buffered saline pH 7.4) for TEM samples was prepared as: 0.876 9 NaCl, 25 mL of 0.2M Na-Cacadylate buffer (Electron Microscopy Sciences Fort Washington, PA), 5 mL 50% glutaraldehyde (Electron Microscopy Sciences Fort Washington, PA), distilled water up to 100 mL and adjust the pH to 7.4. Secondary fixation 18 was done in 1% Osmium tetroxide (Electron Microscopy Sciences Fort Washington, PA). The samples were dehydrated in an acetone gradient (Electron Microscopy Sciences Fort Washington, PA). ,Spurr resin was obtained from the Center for Electron Optics at Michigan State University and was prepared as: 10 g vinylcyclohexene, 6 g diglycidyl ether of polypropylene glycol, 26 g nonenyl succinic anhydride, and 0.4 g of dimethylaminoethanol. Lead stain was prepared as described by Hanaichi et al. (1986) briefly: 0.20 g of calcined lead citrate, 0.15 9 lead nitrate, 0.15 9 lead acetate, 1 g sodium citrate, and 41 mL of distilled water were placed in a 50 mL volumetric flask and mixed well to yield a yellowish, milky solution. Then, 9.0 mL of 1N NaOH was added to the solution and mixed well until the solution became a transparent yellowish color. Two hundred-mesh nickel grids (Electron Microscopy Sciences Fort Washington, PA) were used for mounting ultra-thin sections for post staining and TEM evaluation. Carbon coating of specimens was carried out in vacuum evaporator LADD (Williston, VT). A Phillips CM-10 transmission electron microscope (Hillsboro, OR) was used to examine the specimens. Thin sections of the resin embedded tissue and fresh tissue were mounted on glass slides for examination at the light microscopy level. Molecular biology materials. Primary disruption of rhinosporidial tissue samples were done in a under liquid nitrogen using a mortar and pestle. Secondary disruption of rhinosporidial tissue was performed in 3 mL vacutainer tubes (Becton Dickinson Rutherford, New Jersey) using 500 pm glass beads from the (Microbeads Co. Cleveland, 19 OH). Protein and RNA were digested with Proteinase K and Rnase One respectively (Promega Madison, WI). Nucleic acids were quantified using a GeneQuant DNA/RNA calculator (Amersham Pharmacia Biotech Piscataway, New Jersey). PCR amplifications were performed in a Perkin Elmer (Foster City, CA) GeneAmp® 9700 therrnocycler and the primers used for this study were synthesized using the trityl method in the Macromolecular Structure Facility at Michigan State University. Electrophoresis for DNA separation was performed using 0.7% (cracking gel), 0.8% (genomic DNA), or 1.2% agarose (PCR reactions) Promega (Madison, WI) in TAE buffer. DNA within the gels was stained with 50 pg/mL of ethidium bromide and visualized under ultraviolet light on a Bio-Rad Gel Doc 1000 gel documentation system (Bio—Rad Hercules, CA). A TOPO TA Cloning® Kit from lnvitrogen (Carlsbad, CA) was used to clone all PCR amplicons of interest in this study. Miniprep DNA for sequencing was isolated with the S.N.A.P. Miniprep Kit from lnvitrogen (Carlsbad, CA). ABI Prisms’ Big DyeTM Terminator Cycle Sequencing Ready Reaction Kit (Perkin Elmer Foster City, CA) chemistry was used to facilitate the sequencing. Sequencing reactions were run in a GeneAmp® 9700 therrnocycler and the completed, cleaned up sequencing reactions were sequenced using an ABI 310 genetic analyzer (Perkin Elmer Foster City, CA). Sequences were analyzed using ABI Prism Sequencing Analysis and Sequence Navigator software from Perkin Elmer (Foster City, CA) and Pyhlogenetic Analysis Using Parsimony (PAUP) version 3.1 D.L. Swofford Illinois Natural History Survey (Champaign, Ill). 20 METHODS Collection and storage of tissues containing R. seeberi sporangia and endospores. Biopsied tissue, containing R.seeberi sporangia and endospores, were collected in Sri Lanka by Dr. Sarath N. Arseculeratne from two men with rhinosporidiosis. The tissues were aseptically collected, washed in sterile water, and immediately put on ice and shipped to Dr. Leonel Mendoza at Michigan State University. Prior to the extraction of DNA the tissue samples were washed in sterile deionized water three times for 10 minutes each at room temperature with gentle agitation. Part of the final wash was used to inoculate a saubourauds agar plate to detect possible contaminants. The remaining washes were pooled and centrifuged at 500 g for 10 minutes to collect the free endospores that were released during the washing process. The endospore pellet was resuspended using sterile deionized water and stored at -20°C. A small amount of tissue from each sample was placed onto a glass slide with a drop of water for microscopic examination. Each sample was examined for gross contamination as well as to confirm the presence of R. seeberi in the tissue samples. The two rhinosporidial tissue samples were then dissected into 100 mg blocks and placed in microfuge tubes for storage at -80°C. 21 Transmission electron microscopy studies on rhinosporidial tissues. Preparation of sample blocks: Several 1 mm cubes of infected human nasal tissue containing sporangia and endospores of R. seeberi were brought to room temperature (from -80°C). These samples were placed in 2.5% glutaraldehyde with 0.05M Na-Cacadylate buffered saline (pH 7.4), incubated at room temperature for 2 hours with gentle agitation. This primary fixation was followed by three 0.05M Na-Cacadylate buffered saline washes for 20 minutes each. The sample was then placed into a 1% 0804 in water solution at room temperature for 4 hours with mild agitation. The 0804 post fixation was followed by three 20 minute distilled water washes. Dehydration in acetone was accomplished by using a stepwise. gradient; 25%, 50%, 75%, and two 100% acetone changes for 30 minutes in each step. The final dehydration step was overnight in 100% acetone. The infiltration of the sample was accomplished by using a stepwise gradient. Samples were transferred to 33% followed by 66% spurr resin in acetone solutions for 30 minutes each. The sample was then put into 100% spurr resin for 2 hours, 5 hours, and overnight with resin changes at the end of each time period. The samples were cast in fresh spurr resin in an open mold and put in a 65° C oven for 48 hours. 22 Sectioning blocks: The blocks of interest were examined and then trimmed down for sectioning by using a razor blade and a microtome chuck and chuck holder. Trimmed blocks were placed in the MTX ultra-microtome (RMC Tucson, AZ), faced-up and thin sectioned. Thin sections (750nm) were removed and heat fixed to a glass slide and stained with 1% toludine blue in a 1% sodium borate solution for 1 minute at 55°C. Several light microscopy pictures were taken on a BH-2 Olympus microscope at 40X and 100X. Ultra-thin sections were made using the MTX ultra-microtome to section the sample into 100 nm thick sections that were floated onto 200 mesh Ni grids. The grids with the ultra thin sections were post stained using 3% uranyl acetate for 30 minutes followed by Hanaichi’s lead (Hanaichi et al. 1986) for 3 minutes. After the post staining procedure, the grids were put into a LADD vacuum evaporator and a thin layer of carbon was evaporated onto the surface of the grids. Examination of ultra-thin sectioned material: The grids were examined using a Phillips CM-10 electron microscope. Micrographs of the ultra-thin sectioned material were taken at 11,500X and 52,000X . Molecular studies on rhinosporidial tissues. Genomic DNA isolation from rhinosporidial tissue: Genomic DNA from both rhinosporidial polyps was isolated using the following protocol. After the rhinosporidial tissue samples were collected, 23 washed, and stored as described earlier, DNA extraction was initiated by removing a single 100 mg block of sample tissue from —80°C to a mortar that had been ore-cooled with liquid nitrogen. The rhinosporidial tissue was ground under liquid nitrogen for 20 to 30 minutes until the tissue was reduced to a fine chalk- like powder. Fifty mg of the powder was transferred to a sterile red top 3 mL vacutainer containing 300 pL of sterile glass beads (500 BM in diameter). The remaining powder in the mortar was transferred to a second vacutainer tube containing 300 pL of glass beads. Seven hundred and fifty pL of sterile deionized water was put into each vacutainer to resuspend the powder, tubes were incubated on wet ice for four minutes, vortexed at the highest setting for two minutes, and returned to the wet ice for an additional four minutes. Ten pL of the supernatant from each tube was placed on a microscope slide to visually inspect the samples for endospore disruption. The samples were repeatedly placed on ice and vortexed until greater than 90% disruption of the endospores was obtained. Two hundred and fifty pL aliquots of the supernatant were transferred to 1.5 mL sterile snap top microfuge tubes containing 250 uL of lysis buffer each. The tubes (containing a total volume of 500 uL) were inverted gently 15 to 20 times and proteinase K was added to a final concentration of 250 pg/mL. The tubes were placed in a 55°C water bath for one hour, and then 500 pL of Tris saturated phenol: chloroform: isoamyl (25:24:1) was added to the tubes. All of the tubes were incubated at 25°C with gentle agitation for 15 minutes. After the incubation the tubes were centrifuged at 12,000 g for 15 minutes at room temperature and the aqueous phase was carefully removed 24 from each tube (~300 pL) to a separate, sterile 1.5 mL microfuge tube. Ten pL of 3M sodium acetate (pH 4.6), followed by two volumes of ice cold 100% ethanol was added to each tube and the tubes were gently inverted 5-6 times. The nucleic acids were precipitated overnight at -20°C. The following day the tubes were centrifuged for 15 minutes at 4°C and 12,000 g. The ethanol supernatant was carefully aspirated from each of the tubes and 900 pL of 70% ethanol was carefully added to each tube to wash the pellets. The tubes were centrifuged for 2 minutes at room temperature and the 70% ethanol was aspirated off. The tubes were placed in a dessicator for 15 minutes (no vacuum) to dry the pellets. The pellets were resuspended in 50 pL of sterile deionized water. All of the resuspended nucleic acids from each sample were pooled and put through a second ethanol precipitation as described above to concentrate all of the isolated nucleic acids from each sample in one tube. The pellets were resuspended in 150 ILL of sterile deionized water and stored at -80°C. To confirm the presence of extracted nucleic acids in the samples (extraction protocol described above), 10 pL aliquots of rhinosporidium nucleic acids from each patient sample mixed with 2 pL of 6X loading dye from each tube were loaded into a 0.8% agarose TAE gel with 0.5 pg/mL of ethidium bromide and run at 135 volts for one hour. The remainder of the samples were subjected to an RNase treatment. Ten pL of RNase One, 400 pL of sterile deionized water, and 50 DL of the RNase One 10X buffer was added to the 150 uL of each sample. The tubes were gently inverted 10-12 times and placed in a 37°C water bath for one hour and fifteen minutes. The RNase treatment was 25 followed by a phenol chloroform extraction and ethanol precipitation as described earlier. The pellets were resuspended in 150 pL of sterile deionized water and stored at -80°C. Ten uL aliquots of the RNase treated rhinosporidium DNA from each tube were mixed with 2 pL of 6X loading dye for each sample and loaded into a 0.8% agarose TAE gel with 0.5 pg/mL of ethidium bromide and run at 135 volts for one hour. The ethidium bromide stained agarose gels were visualized on a Gel Doc 1000 digital image capture system from Bio-Rad (Hercules, CA USA). Part of each sample was diluted with sterile deionized water to a working dilution of 100ng/pL for PCR and stored at -20°C, and the remainder of the original stock was stored at -80°C. PCR amplification of the 18S serNA: Due to the limited quantity of R. seeberi DNA available for our experiments, DNA from the oomycete Pythium insidiosum was used to standardize the 188 serNA PCR experiments. DNA from P. insidiosum was extracted, purified, and diluted to a working concentration of 100 nglpL following the same protocol as described eariier. Once the 188 serNA PCR was standardized using P. insidiosum DNA, PCR experiments using R. seeberi as sample template were performed. In these experiments P. insidiosum DNA was used as a positive control template. Twenty-five pL PCR reactions were set up on wet ice, under a biological safety hood in 0.2 mL MicroAmp ® reaction tubes from (Perkin Elmer, Foster City, CA). The sample reactions consisted of: 2.5 pL of GeneAmp® 10X PCR buffer II (no MgClz) (Perkin Elmer, Foster City, CA), 1.5 pL of 25mM MgClz 26 solution (Perkin Elmer, Foster City, CA), 2 pL of 10 mM dNTP mix (Perkin Elmer, Foster City, CA), 5 pmoles of a forward primer (table 2), 5 pmoles of a reverse primer (table 3), 0.25 pL of AmpliTaq GoldTM(Perkin Elmer, Foster City, CA), 500 ng of DNA template, and sterile deionized water to Q. S. the reaction volume to 25 BL. A primer map (figure 2) shows the sites where the forward and reverse primers that were used in this study are located. A reagent blank (no template) as well as a positive control (500 ng P. insidiosum DNA), were run with the samples during each PCR experiment. The PCR thermo-cycling parameters were as follows: 10 minutes at 95°C followed by 40 cycles of 1 minute at 95°C, 2 minutes at 50°C, and 3 minutes at 72°C followed by 1 cycle of 1 minute at 95°C, 2 minutes at 50°C, and 10 minutes at 72°C. PCR reactions were run out on a 1.2% agarose (Promega) TAE gel containing 0.5 pg/ mL of ethidium bromide, and documented on a Gel Doc 1000 system from Bio-Rad (Hercules, CA USA). 27 Table 2: Abbreviations and sequences of 18S serNA forward primers. Forward Primer Sequence 5’-)3’ Reference Primers NS 1 gTA gTC ATA TgC TTg TCT C NS 3 gCA AgT CTg ng CCA gCA gCC NS 5 AAC TTA Aag gAA TTg ACg gAA 9 N87 gAg gCA ATA ACA ggT CTg TgA TgC Table 3: Abbreviations and sequences of18$ serNA reverse primers. Reverse Primer Sequences 5’93’ . Reference Primers NS 2 990 T90 T99 CAC CAg ACT TgC NS 4 CTT CCg TCA ATT CCT TTA Ag NS 6 gCA TCA CAg ACC TgT TAT TgC CTC NS 8 TCC gCA ggT TCA CCT ACg gA NS 8 DE. TCC gCA ggT TCA CCT(A) AngA 28 18S PRIMER MAP 500 bp L—_I N83 NS 5 —> —> Figure 2: Primer map of the 18S rDNA molecule with the locations of N81, 2, 3, 4, 5, 6, 7, and N88 degenerate primers (Gargas and DePreist1996). Cloning of the 18S serNA amplicon of R. seeberi: The PCR protocol outlined above was used to amplify a 1791 b.p. region of R. seeberi’s 18S serNA using the primers NS1 (Gargas and DePriest 1996) and NS8 degenerate (as per lssakainen 1997). The TOPO TA cloning kit (Invitrogen, Carlsbad, California) was used to clone the amplified fragments. The amplicons were cloned into the pCR 2.1 vector as per the manufacturers instructions. Briefly, twelve micro-liters of the PCR reaction to be cloned was electrophoresed into a 1.2% agarose gel as described eariier to visually confirm that the amplicon of interest was present and that there were no other amplicons present. A 0.25mL MicroAmp tube was used to set up a 5 (IL cloning reaction containing in order: 3uL of fresh PCR product, 1 pL of sterile deionized water, and 1 pL of pCR®-2.1 TOPO® vector. The cloning reaction was mixed gently 29 and incubated at room temperature for 5 minutes. Immediately following the 5 minute incubation 1 pL of the 6X TOPO® Cloning Stop Solution was added and the reaction was mixed gently for ~10 seconds. The cloning reaction was then immediately placed on ice. Two tubes of TOP10 One ShotTM competent cells were thawed out and the cloning reaction was split into 2pL and 3pL portions that were added to each of the tubes of TOP10 cells and mixed gently. The transformation reactions were incubated on ice for 30 minutes and then were heat shocked for 30 seconds at 42°C without Shaking. After heat shocking the cells, the reactions were immediately transferred to ice and 250pL of room temperature SOC medium lnvitrogen (Carlsbad, CA USA) was added to each tube of the transformation reactions. The transformed cells in SOC were shaken horizontally at 37°C for one hour (for kanamycin selection) and then placed on ice. Six 100mm LB agar plates with 50pg/mL of kanamycin Sigma (St. Louis, MO USA) were warmed up and 40pL of a 40pglmL X-gal solution Promega (Madison, WI USA) was spread evenly onto each plate and the plates were placed at 37°C to dry. Each transformation reaction was plated onto three separate LBK plates in 100pL, 50pL, and 10pL volumes. All of the plated material was incubated overnight at 37°C. The plates were removed to 4°C for 4 hours to allow the transfomrants with no inserts to turn dark blue. Isolated white colonies were selected from all six plates, and a dark blue colony was selected from each plate for use as a negative control. The putative positive clones and the negative controls were sub-cultured onto LBK agar plates which were divided 3O into 4 equal areas (4 clones/plate). The sub-cultures were incubated overnight at 37°C and put at 4°C until further analysis. Analysis of putative positive clones: To confirm that the sub-cultured putative positive clones have been transformed by a pCR 2.1 vector which contains an insert, all of the sub-cultured clones were analyzed by cracking gel (Maniatis et al. 1982). A cracking gel is a technique used to rapidly disrupt bacterial colonies to test the size of their plasmids. Briefly, using a sterile toothpick part of each sub-cultured colony is transferred to a separate labeled 0.65mL microfuge tube containing 50p.L of a sterile 10mM EDTA (pH 8.0) solution. Fifty micro-liters of freshly made cracking buffer (page 14) was added to each tube and the tubes were vortexed for thirty seconds. The tubes were incubated at 70°C for 15 minutes and then were cooled to room temperature. One and a half micro-liters of 4M KCI and 0.5pL of 0.4% bromophenol blue were added to each tube and the tubes were all vortexed for 30 seconds. The tubes were transferred to ice for 5 minutes and were spun down at 12,0009 for 3 minutes at 4°C to remove bacterial debris. Fifty micro-liters of the supernatant from each tube was loaded into each well of a 0.7% agarose in TAE gel (containing 50pg/mL of ethidium bromide) and run until the bromophenol blue dye front had migrated % of the way down the gel. The cracking gels were analyzed on a Gel Doc 1000 gel documentation system Bio-Rad (Hercules, CA USA). Archival storage of clones: 31 After confirming the presence of plasmids with inserts in the positive Clones, glycerol stocks were made of each Clone for archival storage at -80°C. Briefly, sub-cultured positive clones were streaked onto LB kanamycin plates (Song/mL) for isolation of single colonies, and were grown overnight at 37°C. A single colony representing each clone was transferred to a separate sterile tube containing 2 mL of LB kanamycin (SOpglmL) broth and grown at 37°C with shaking until the culture reached mid-log phase. Eight hundred and fifty micro- liters of each clone’s culture was mixed with 150 pL of sterile glycerol and stored at -80°C in a 1.5 mL cryovial. Isolation of plasmid DNA from E. coli: Plasmid DNA from clones of interest were purified using the S.N.A.P. Miniprep Kit from lnvitrogen (Cadsbad, CA USA). The plasmid DNA was purified by following the manufacture’s instructions. Briefly, a 3mL culture of a pure clone was grown overnight at 37°C in LB kanamycin broth (50pg/mL). The cells were pelleted at 5009 for 10 minutes and the supernatant was removed. The cell pellet was resuspended using 150 pL of resuspension buffer and gently pipetting up and down. One hundred and fifty micro-liters of lysis buffer was added and the tube was inverted gently 5-6 times. The tubes were then incubated at room temperature for 3 minutes. One hundred and fifty micro-liters of ice-cold precipitation buffer was added and the tubes were inverted gently 6-8 times. The mixture was spun in a microcentrifuge for 5 minutes at 14,0009. After centrifugation, the supernatant was removed to a sterile 1.5 mL microcentrifuge tube and the gelatinous pellet was discarded. Six hundred 32 micro-liters of binding buffer was added to the supernatant and the tube was inverted 5-6 times. The entire solution was then pipetted into a S.N.A.P. miniprep column/collection tube and centrifuged at 3,0009 for 30 seconds. The column flow thrOugh was discarded and 500 pL of wash buffer was added to the column. The column/collection tube was centrifuged for 30 seconds at 3,0009 and the column flow through was discarded. Nine hundred micro-liters of the 1X final wash was added to the column, and the column/collection tube was spun down for 30 seconds at 3,0009. The column flow through was discarded and the column/collection tube was spun down for 1 minute at 3,0009 to dry the resin. The miniprep column was placed into a fresh sterile 1.5 mL microcentrifuge tube and 60uL of sterile deionized water was added to the resin in the column. The . hydrated column was incubated for 5 minutes at room temperature and then the tube was spun in a microcentrifuge at maximum speed for 30 seconds. The eluted plasmid DNA was stored at -20°C and the column was discarded. Sequencing the miniprep plasmid DNA: Sequencing reactions were prepared using the ABI PRISMTM BigDye'l'M Terminator Cycle Sequencing Ready Reaction Kit with AmpliTaq® DNA Polymerase, FS from Perkin Elmer Applied Biosystems (Foster City, CA USA). A modification of the standard 20 pL reaction which is described by the manufacturer was used. The modified sequencing reactions were assembled in 0.25mL MicroAmp® tubes on ice from Perkin Elmer as described in table 4 below (Foster City, CA USA). In addition to the forward and reverse primers listed in tables 2 and 3 -21 M13 (fonrvard) universal primer and -24 M13 (reverse) 33 universal primer Promega (Madison, WI USA) were used in sequencing reactions. Sequencing reactions were mixed briefly and spun down in a microcentrifuge. A GeneAmp® 9700 thermal cycler was programmed to repeat the following thermal profile for 25 cycles: rapid thermal ramp to 96°C, hold at 96°C for 10 seconds, rapid thermal ramp to 50°C, hold at 50°C for 5 seconds, rapid thermal ramp to 60°C, and hold for 4 minutes at 60 °C. After the 25th cycle the reactions were cooled to 4°C and held until they were purified. 34 Table 4: ABI PrismTM Big DyeTM Terminator Cycle Sequencing reaction. Terminator Ready Reaction Mix 4.0pL Template (Miniprep DNA) 200-500ng Primer 3.2pmoles Deionized water q.s. TOTAL VOLUME 10pL 35 The sequencing reactions were “cleaned up” using an ethanol precipitation to remove excess dye terminators from the reaction. Briefly, the 10pL reactions were spun down and the entire reaction was added to a Genetic Analyzer Sample Tube (0.5 mL) Perkin Elmer (Foster City, CA USA) containing 32pL of room temperature 95% ethanol and 69L of sterile deionized water. The tube was vortexed briefly and the mixture was allowed to precipitate at room temperature for 15 minutes. After the precipitation the tube was spun down in a microcentrifuge at full speed for 30 minutes. The supernatant was carefully aspirated and discarded, taking care not to disturb the small pellet. The open tube was placed in a vacuum centrifuge to dry for 5 minutes. The dried pellet was resuspended in 18p.L of TSR (template suppression reagent from Perkin Elmer (Foster City, CA USA)). The sample was briefly vortexed and spun before it was heated for 2 minutes at 95°C to denature any double stranded DNA. Immediately following the 95°C incubation the tube was transferred to ice for 10 minutes. The sample was vortexed and spun down again and a Genetic Analyzer Septa for 0.5 mL Sample Tubes Perkin Elmer (Foster City, CA USA) was inserted into the tube. The samples were loaded into an ABI Prism 310 Genetic Analyzer Perkin Elmer (Foster City, CA USA) and run on a 61 cm x 50 um internally uncoated capillary using POP-6 polymer and 310 Genetic Analyzer Buffer with EDTA Perkin Elmer (Foster City, CA USA). The samples were run using the Seq POP6 (1mL) E run module, the DT POP6 {BD Set-Any Primer} 36 dye set/primer file, and the raw data was analyzed using a matrix that was constructed on the same 310 for analyzing BigDyeTM terminator chemistry. Analysis, alignment, and editing of the nucleic acid sequences: Sequence data from the ABI PrismTM 310 Genetic Analyzer Perkin Elmer (Foster City, CA USA) was aligned, edited, and contigs were constructed using Perkin Elmer’s (Foster City, CA USA) Sequencing Analysis and Sequence Navigator software. Pyhlogenetic Analysis: The 188 serNA sequence of R. seeberi was aligned by visual inspection with sequences previously analyzed by (Spanggaard et al. 1996 and Ragan et al. 1996) with Sequence Navigator Perkin Elmer (Foster City, CA USA). Regions of ambiguous alignment were excluded from phylogenetic analysis, and gaps were treated as missing data. Phylogenetic analysis was by a distance method (neighbor joining with Kimura’s 3-parameter multiple-hit correctionXSpanggaard et al. 1996) and parsimony. Support for internal branches was assessed by using 1,000 bootstrapped data sets. Parsimony analyses were conducted with the computer software program PAUP (PAUP: Phylogenetic Analysis Using Parsimony, version 3.1; D. L. Swofford, Illinois Natural History Survey, Champaign, Ill.). Parsimony analysis employed 1,000 heuristic searches with randomized orders of taxon entry to increase the chance of finding the shortest trees. Trees were drawn with stramenopiles and alveolates as the outgroups, based on previous phylogenetic studies. (Kerk eta/1995. , Ragan et al. 1996, and Spanggaard et al. 1996) 37 Chapter 3 RESULTS Histopathology and transmission electron microscopy (TEM). Several light microscopy photographs were taken to document the presence of R. seeberi in the tissue samples used for DNA analysis. The micrographs reveal the presence of many juvenile and intermediate sporangia as well as mature sporangia containing immature and mature endospores. Histopathologically, a cell-mediated immune response composed of macrophages in the process of phagocytizing endospores and sporangia, giant cells, and plasma cells was observed. lntennediate sporangia with Clearly visible nuclei and prominent nucleoli were also observed. Transmission electron microscopic examination of the rhinosporidial tissue revealed details of the host parasite relationship. Several “dead” intermediate sporangia which had been killed by the host’s immune response were observed. These “dead” sporangia possessed several unique features. In contrast to a healthy intermediate sporangium, the “dead” sporangiums’ cytoplasm appeared to be homogeneous with no apparent organelles or membrane bound structures. It was also noted that the cell wall in these sporangia appeared thicker and less dense than a healthy cell wall, it also was 38 noted that the outer edge of this cell wall was very uneven and possessed fibrous clumps on its surface as well as in the surrounding inflammatory tissue. The second feature that was observed in healthy sporangia was the presence of mitochondria clusters (usually 4-7). The mitochondria of R. seeben' were only observed in intermediate sporangia. They were approximately 5009m in diameter and possessed flat mitochondrial cristae. Checking for the presence and quality of genomic DNA from Rhinosporidium seeberi. To investigate the quality of the genomic DNA samples from Sri Lanka, the isolated DNA was electrophoresed into a 0.8% agarose gel (see figure 3). In figure 3 the outside lanes of the gel are a high molecular weight marker (23.1 Kb - 564bp) from Bio-Rad (Hercules, CA USA). Lanes 2 and 3 contain 10th of a 1:10 dilution of R.seeberi nucleic acids and 10pL of undiluted sample respectively. It was observed that the presence of a faint 23.1 Kb band of genomic DNA was present in lanes 2 and 3 which is trailed by a very heavy smear of degraded DNA from 23.1 Kb down. The vast majority of the genomic DNA has been partially degraded by endonucleases prior to and during the isolation of the sample. In addition, there are 5-6 faint, but observable bands of less than 2,028 bp visible in lane 2. 39 Figure 3: An 0.8% agarose gel showing a 1:10 dilution and undiluted nucleic acids of R. seeberi prior to RNase treatments in lanes 2 and 3 respectively. 40 23.1 Kb Figure 4: This figure shows a 0.8% agarose gel showing R. seeberi DNA after three RNase digests. Lanes 2, 3 and 4 contain 59L of sample DNA from separate RNase treatments. 41 Verification of RNase digestion. The quality of nucleic acid sample was analyzed after three RNase digestions and phenol extractions. Figure 4 shows lanes 2,3,and 4 with 5pL of R. seeberi genomic DNA. A high molecular weight marker from Bio-Rad (Hercules, CA USA) was run in lane 1. It was observed that after the RNase treatments a weakened banding pattern at the end of the gel remained in all three of the RNase treated samples (lanes 2,3 and 4). The 23.1 Kb band that was present in the untreated samples (see figure 3) was no longer visible. DNA that remained in the samples was degraded and produced a heavy smear from 23.1 Kb to less than 500 bp in lanes 2,3 and 4 (see figure 4). PCR amplification of the 188 rDNA from Rhinosporidium seeberi. To optimize the PCR conditions for amplifying the 188 rDNA from R. seeberi numerous samples were run using a variety of different PCR conditions. It was determined that the optimal conditions were: 2.5 pL of GeneAmp® 10X PCR buffer II (no MQCIz) (Perkin Elmer, Foster City, CA), 1.5 pL of 25mM MQCIz solution (Perkin Elmer, Foster City, CA), 2 pL of 10 mM dNTP mix (Perkin Elmer, Foster City, CA), 5 pmoles of N81, 5 pmoles of NS8D, 0.25 pL of AmpliTaq GoldTM(Perkin Elmer, Foster City, CA), 500 ng of R. seeberi DNA template, and sterile deionized water to Q. S. the reaction volume to 25 pL. The optimal PCR thermo-cycling parameters were as follows: 10 minutes at 95°C followed by 40 cycles of 1 minute at 95°C, 2 minutes at 50°C, and 3 minutes at 72°C followed by 1 cycle of 1 minute at 95°C, 2 minutes at 50°C, and 10 minutes at 72°C. These conditions produced a single amplicon of 1791 base pairs. Aliquots of the PCR reactions that were intended for cloning purposes were run on a 1.2% 42 agarose gel to verify the presence of an amplicon of the expected molecular mass (see figure 5). In figure 5 lanes 4-7 were loaded with 12pL aliquots from each of the 259L PCR reactions that had been run under the optimal conditions described above but, the amount of R. seeberi template in each was varied: 100ng (lane 4), 300n9 (lane 5), 500n9 (lane 6), 700ng (lane 7). Lanes 2 and 3 were a P. insidiosum (500n9) positive control and a reagent blank (no template) respectively. It was observed that the PCR reaction loaded into lane 6 (500 ng of R. seeberi template) yielded the best amplicon. Therefore, the remainder of that PCR reaction was used as the insert DNA for the subsequent cloning reaction. 43 1234567 Figure 5: This figure shows a 1.2% agarose gel used to document the presence of the amplicon of predicted size prior to cloning. 44 Cloning of the 1791 base pair amplicon, and cracking gel analysis of putative positive clones. After the 1791 base pair amplicon was cloned into TOP10 cells with the pCR 2.1 vector from lnvitrogen (Carlsbad, CA USA), putative positive (white) colonies were picked for further analysis. It was observed that the plates with 100pL, 50pL, and 10pL volumes of transformed cells all had 50-250 putative positive colonies each. Fifty-four putative positive clones were selected randomly from the three plates as well as 5 blue colonies for negative controls. All 54 “positive” clones and the 5 negative control clones were further analyzed by cracking gels (see figures 6-9). Cracking gel #1 (clones 1-15) figure 6 and cracking gel #2 (clones 16-30) figure 7 were run at 110V for 3 hours. The negative controls (pCR2.1 vector with no insert) were run in lanes 1 and 16 of figures 6 and 7 respectively. In both figures, the plasmid DNA from the negative controls was significantly smaller than that of the putative positive clones. Gels #1 and 2 were run to long and the DNA bands present in the gel are not very well defined. Cracking gel #3 (clones 30-44) figure 8 and cracking gel #4 (clones 45-49, and 54-63) figure 9 were run at 100V for 2 hours. The negative controls were run in lanes 33 for cracking gel #3,and 45 and 61 in cracking gel #4. In both gels the negative controls were significantly lower than all of the putative positive clones. It was found that running the cracking gels at 100V instead of 110V and decreasing the length of the run to 2 hours gave better separation and definition of the plasmid DNA. Due to the poor definition of bands in cracking gels #1 and 2, clone 30 was run in cracking gel #3 to confirm 45 the results of cracking gel #2. It was observed that clone 30’s plasmid DNA was significantly larger than the negative control plasmid DNA in both cracking gels # 2 and #3. 46 Figure 6: Cracking gel #1, (clones 1-15) was run for 3 hours at 110V. Lane #1 contains the negative control (pCR2.1 vector with no insert). 47 ., , urn-9‘. I. Figure 7: Cracking gel #2 (clones 16-30) was run for 3 hours at 110V. Lane 16 is the negative control (pCR 2.1 vector with no insert). 48 * 44434241403938 37 3635343332 3130 Figure 8: Cracking gel #3 (clones 30-44) was run for 2 hours at 100V. Lane 33 contains a negative control (pCR 2.1 vector with no insert). Lane 30 contains clone #30 which was repeated to confirm the results of cracking gel #2, clone 30. 49 636261 m$5857 5655544948 474645 Figure 9: Cracking gel #4 (clones 45-49 and 54-63) was run for 2 hours at 100V. Lanes 45 and 61 contain negative controls (pCR 2.1 vector which contains no insert). 50 Sequencing of Rhinosporidium seeberi’s 188 rDNA. After cracking gels were run to confirm the presence of an insert, nineteen clones were randomly selected to be sequenced using N85 and NS8 degenerate primers (tables 2 and 3), to get 529 base pairs of complete overlap (for each clone) within the 18S molecule. After further analysis of the clones it was found that the 529 base pair fragment began at base 1,196 of the complete 1,791 base pair amplicon of R. seeberi’s 18S rDNA, and went up to and included base 1,724 (figure 10). After editing the overiapping sequences for each clone, all 19 of the 529 base pair fragments (one fragment per clone) were put through a simple BLAST search using GenBank. The results of the search indicated that all 19 fragments were part of an 188 rRNA gene that was 98-96% homologous to an identical region of an 18S rRNA gene from Dermocystidium sp. It was also observed that there were no human, animal, or fungal sequences listed in the search results. When all 19 of the cloned fragments were aligned and compared to one another several polymorphic sites were found (figure 10). It was found that of the 19 screened clones, there were 8 different genotypes due to single point mutations at 7 different sites. Within the 529 base pair region; 10 of the Clones were identical (2, 3, 6, 7, 9, 11, 14, 17, 20,and 22) and were not polymorphic at any of the 7 sites, Clones 4, 5, 8, 12, and 21 all had unique single point mutations, and clones 10 and 15 as well as clones 18 and 19 were identical pairs that each had the same single point mutation. The 529 base pair sequence of Clone RS #2 is shown in figure 10 (Rs #2 is identical to Rs 3, 6, 7, 9, 11, 14, 17, 20, and 22 within this region) with the 7 polymorphic sites highlighted. 51 The polymorphic sites are as follows: base 1326, base 1343, base 1377, base 1423, base 1468, base 1505,and base 1711. Clones Rs #2, Rs #4, and Rs#20 were Chosen to be entirely sequenced. Primers NS1,_N82, N83, N84, N85, N86, N87, N88 degenerate (tables 2 and 3), and the universal primers M13 -21 and M13 -24 lnvitrogen (Carlsbad, CA USA) were all used on all 3 clones to get as much overlapping sequence as possible. All three clones had at least 2 sequencing reactions covering every base in the entire sequence. On all three clones at least 90% of the entire sequence had at least 3 sequencing reactions covering each base. After the sequences for each clone were edited against themselves, they were put through a simple BLAST search using GenBank. The results of the search were identical to those of the previous search using the 529 base pair fragment of R. seeberi’s 18S rDNA. Once again the 18S rDNA from Dermocystidium sp. was the closest match. The 3 complete 188 sequences were aligned and compared to each other. It was found that of the 3 clones that were entirely sequenced there were 3 separate genotypes. The differences between the clones were all due to point mutations at the following sites: base 397, base 535, base 1343,and base 1777. The entire sequences of all 3 clones and the locations of the polymorphic Sites are shown in figure 11. 52 1205 1245 Rs#Z ACGGGGAAAC TCACCAGGTC CAGACATAGT AAGGATTGAC AGATTGAGAG 1255 1295 CTCTTTCTTG ATTCTATGGG TGGTGGTGCA TGGCCGTTCT TAGTTGGTGG 1305 1345 AGTGATTTGT CTGGTTAATT CCGTTAACGA §CGAGACCTT AACCTGCEAA 1355 1395 ATAGTTGCGT GATTTTCGAA TCATTTTAAC TQCTTAGAGG GACTATTGGT 1405 1445 GTTTAACCAA TGGAAGTTTG AGGCAATSAC AGGTCTGTGA TGCCCTTAGA 1455 1495 TGTTCTGGGC CGCACGCGCG CTECACTGAT AAAGGCAACG AGTTTATCAC 1505 1545 CTGTACCGGg AGGTATGGGT AATCTTTTCA.AACTTTATCG TGCTGGGGAT 1555 1595 AGATCTTTGC AATTTTCGAT CTTAAACGAG GAATTCCTAG TAAGCGCAAG 1605 1645 TCATCAGCTT GCGTTGATTA CGTCCCTGCC CTTTGTACAC ACCGCCCGTC 1655 1695 GCTACTACCG ATTGAATGGT TTAGTGAGGT CTTCGGATTG GCGCTTTGCA 1705 1724 GCTGGCGACA GCAGTflGGAT GCCGAAAAG Figure 10: The figure depicts a 529 base pair fragment (bases 1196-1724) of the complete 1,791 amplicon from R. seeberi’s 18S rDNA molecule. This fragment is flanked by N85 and N88 degenerate primers, and was utilized to screen 19 clones that were confirmed to have inserts by cracking gel analysis. All nineteen 529 base pair fragments were put through a simple BLAST search using GenBank. The search results indicated that all 19 fragments were part of an 188 rRNA gene that was 98—96% homologous to the same region of an 188 rRNA gene from Dermocystidium sp. Of the 19 clones that were screened there were 8 different genotypes due to single point mutations at 7 different sites. Within the 529 base pair region; 10 of the clones were identical (2, 3, 6, 7, 9, 11, 14, 17, 20,and 22), clones 4, 5, 8, 12, and 21 were all unique, and clones 10 and 15 as well as clones 18 and 19 were identical pairs. The polymorphic sites are highlighted and are as follows: 1326, 1343, 1377, 1423, 1468, 1505,and 1711. 53 Figure 11: Rhinosporidium seeberi's 188 serNA (1791 base pairs). This figure shows the entire 1,791 base pair sequence of R. seeberi’s 188 rDNA for Clones Rs #2, 4, and 20. The polymorphic sites are highlighted and are as follows: base 397, base 535, base 1343, and base 1777. #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #20 #2 #4 #20 10 GTAGTCATAT GTAGTCATAT GTAGTCATAT 70 CTGTGAAACT CTGTGAAACT CTGTGAAACT 130 TTGGATAACC TTGGATAACC TTGGATAACC 190 ATGTATTTAT ATGTATTTAT ATGTATTTAT 250 TAATAACTTT TAATAACTTT TAATAACTTT 310 ATCAACTTTC ATCAACTTTC ATCAACTTTC 370 GGTTCGATTC GGTTCGATTC GGTTCGATTC 430 GCGCAAATTA GCGCAAATTA GCGCAAATTA 490 TTAAAGTCTT TTAAAGTCTT TTAAAGTCTT 550 GCAAGTCTGG GCAAGTCTGG GCAAGTCTGG 610 GCAGTTAAAA GCAGTTAAAA GCAGTTAAAA 670 TTGTACCGAG TTGTACCGAG TTGTACCGAG 730 GTGCGTCCGT GTGCGTCCGT GTGCGTCCGT 790 ATTGCTTGAA ATTGCTTGAA ATTGCTTGAA GCTTGTCTCA GCTTGTCTCA GCTTGTCTCA GCGAATGGCT GCGAATGGCT GCGAATGGCT GTAGTAATTC GTAGTAATTC GTAGTAATTC TAGATAAAAA TAGATAAAAA TAGATAAAAA GCGAATCGTA GCGAATCGTA GCGAATCGTA GATGGTAAGG GATGGTAAGG GATGGTAAGG CGGAGAGGGA CGGAGAGGGA CGGAGAGGGA CCCAATCCTG CCCAATCCTG CCCAATCCTG GTAATTGGAA GTAATTGGAA GTAATTGGAA TGCCAGCAGC TGCCAGCAGC TGCCAGCAGC AGCTCGTAGT AGCTCGTAGT AGCTCGTAGT CCAGCCAATC CCAGCCAATC CCAGCCAATC CTTATTTCGG CTTATTTCGG CTTATTTCGG TATTTCAGCA TATTTCAGCA TATTTCAGCA AAGATTAAGC AAGATTAAGC AAGATTAAGC CATTAAATCA CATTAAATCA CATTAAATCA TAGAGCTAAT TAGAGCTAAT TAGAGCTAAT ACCAATGCGG ACCAATGCGG ACCAATGCGG TGGCTTCGTG TGGCTTCGTG TGGCTTCGTG TATTGGCTTA TATTGGCTTA TATTGGCTTA GCCTGAGAAA GCCTGAGAAA GCCTGAGAAA ACACAGGGAG ACACAGGGAG ACACAGGGAG TGAGTACAAT TGAGTACAAT TGAGTACAAT CGCGGTAATT CGCGGTAATT CGCGGTAATT TGGATTTTGG TGGATTTTGG TGGATTTTGG CTTTCGCTTC CTTTCGCTTC CTTTCGCTTC GACTTTTACT GACTTTTACT GACTTTTACT TGGAATAATG TGGAATAATG TGGAATAATG 55 CATGCATGTC CATGCATGTC CATGCATGTC GTTATAGTTT GTTATAGTTT GTTATAGTTT ACATGCTAAA ACATGCTAAA ACATGCTAAA ATCTTTTGGG ATCTTTTGGG ATCTTTTGGG CTGACGATGA CTGACGATGA CTGACGATGA CCATGGTTAC CCATGGTTAC CCATGGTTAC CGGCTACCAC CGGCTACCAC CGGCTAICAC GTAGTGACAA GTAGTGACAA GTAGTGACAA TTAAACCCCT TTAAACCCCT TTAAACCCCT CCAGCTCCAA CCAGCTCCAA CCAGCTCCAA GATTTTGGTC GATTTTGGTC GATTTTGGTC TCGAAAGCGG TCGAAAGCGG TCGAAAGCGG GTGAAAAAAT GTGAAAAAAT GTGAAAAAAT GAATAGGACA GAATAGGACA GAATAGGACA TAAGTATAAA TAAGTATAAA TAAGTATAAA ATTTGATAGT ATTTGATAGT ATTTGATAGT AATCCCGACT AATCCCGACT AATCCCGACT TCCGGTTCTT TCCGGTTCTT TCCGGTTCTT TTCATTCAAA TTCATTCAAA TTCATTCAAA AACGGGTAAC AACGGGTAAC AACGGGTAAC ATCTAAGGAA ATCTAAGGAA ATCTAAGGAA AAAATAACAA AAAATAACAA AAAATAACAA TAACGAGGAA TAACGAGGAA TAACGAGGAA TAGCGTATAT TAGCGTATAT TAGCGTATAT GGTTGGTCCG GGTTGGTCCG GGTTGGTCCG CGTGTGCGCT CGTGTGCGCT CGTGTGCGCT TAGAGTGTTC TAGAGTGTTC TAGAGTGTTC TTGGTTCTAT TTGGTTCTAT TTGGTTCTAT 60 CAAATCTATA CAAATCTATA CAAATCTATA 120 ACCTTACTAC ACCTTACTAC ACCTTACTAC 180 TCTGGAAGGG TCTGGAAGGG TCTGGAAGGG 240 TGGTGATTCA TGGTGATTCA TGGTGATTCA 300 TTTCTGCCCT TTTCTGCCCT TTTCTGCCCT 360 GGAGAATTAG GGAGAATTAG GGAGAATTAG 420 GGCAGCAGGC GGCAGCAGGC GGCAGCAGGC 480 TACAGGGCTT TACAGGGCTT TACAGGGCTT 54o CAATTGGAGG CAATgGGAGG CAATTGGAGG soc TAAAGTTGTT TAAAGTTGTT TAAAGTTGTT 660 CCGCAAGGTG CCGCAAGGTG CCGCAAGGTG 720 TAACTGTCGT TAACTGTCGT TAACTGTCGT 780 AAAGCAGGCG AAAGCAGGCG AAAGCAGGCG e40 TTTGTTGGTT TTTGTTGGTT TTTGTTGGTT #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 #2 #4 #20 850 TCTAGGACCA TCTAGGACCA TCTAGGACCA 910 GGTGAAATTC GGTGAAATTC GGTGAAATTC 970 TCATTAATCA TCATTAATCA TCATTAATCA 1030 CCATAAACTA CCATAAACTA CCATAAACTA 1090 ATGAGAAATC ATGAGAAATC ATGAGAAATC 1150 GAATTGACGG GAATTGACGG GAATTGACGG 1210 GAAACTCACC GAAACTCACC GAAACTCACC 1270 ATGGGTGGTG ATGGGTGGTG ATGGGTGGTG 1330 AACGAACGAG AACGAACGAG AACGAACGAG 1390 AGAGGGACTA AGAGGGACTA AGAGGGACTA 1450 TTAGATGTTC TTAGATGTTC TTAGATGTTC 1510 CCGGAAGGTA CCGGAAGGTA CCGGAAGGTA 1570 TCGATCTTAA TCGATCTTAA TCGATCTTAA 1630 CTGCCCTTTG CTGCCCTTTG CTGCCCTTTG AAGTAATGAT AAGTAATGAT AAGTAATGAT TTGGATTTAT TTGGATTTAT TTGGATTTAT AGAACGAAAG AGAACGAAAG AGAACGAAAG TGCCGACTAG TGCCGACTAG TGCCGACTAG AAAGTCTTTG AAAGTCTTTG AAAGTCTTTG AAGGGCACCA AAGGGCACCA AAGGGCACCA AGGTCCAGAC AGGTCCAGAC AGGTCCAGAC GTGCATGGCC GTGCATGGCC GTGCATGGCC ACCTTAACCT ACCTTAACCT ACCTTAACCT TTGGTGTTTA TTGGTGTTTA TTGGTGTTTA TGGGCCGCAC TGGGCCGCAC TGGGCCGCAC TGGGTAATCT TGGGTAATCT TGGGTAATCT ACGAGGAATT ACGAGGAATT ACGAGGAATT TACACACCGC TACACACCGC TACACACCGC TAATAGGGAT TAATAGGGAT TAATAGGGAT GAAAGACTAA GAAAGACTAA GAAAGACTAA TTAGGGGATC TTAGGGGATC TTAGGGGATC GGATTGGTAG GGATTGGTAG GGATTGGTAG GGTTCCGGGG GGTTCCGGGG GGTTCCGGGG CCAGGAGTGG CCAGGAGTGG CCAGGAGTGG ATAGTAAGGA ATAGTAAGGA ATAGTAAGGA GTTCTTAGTT GTTCTTAGTT GTTCTTAGTT GCTAAATAGT GCgAAATAGT GCTAAATAGT ACCAATGGAA ACCAATGGAA ACCAATGGAA GCGCGCTACA GCGCGCTACA GCGCGCTACA TTTCAAACTT TTTCAAACTT TTTCAAACTT CCTAGTAAGC CCTAGTAAGC CCTAGTAAGC CCGTCGCTAC CCGTCGCTAC CCGTCGCTAC 56 AGTTGGGGGC AGTTGGGGGC AGTTGGGGGC CTTCTGCGAA CTTCTGCGAA CTTCTGCGAA GAAGATGATC GAAGATGATC GAAGATGATC ATGTTAATTC ATGTTAATTC ATGTTAATTC GGAGTATGGT GGAGTATGGT GGAGTATGGT AGCCTGCGGC AGCCTGCGGC AGCCTGCGGC TTGACAGATT TTGACAGATT TTGACAGATT GGTGGAGTGA GGTGGAGTGA GGTGGAGTGA TGCGTGATTT TGCGTGATTT TGCGTGATTT GTTTGAGGCA GTTTGAGGCA GTTTGAGGCA CTGATAAAGG CTGATAAAGG CTGATAAAGG TATCGTGCTG TATCGTGCTG TATCGTGCTG GCAAGTCATC GCAAGTCATC GCAAGTCATC TACCGATTGA TACCGATTGA TACCGATTGA ATTAGTATTT ATTAGTATTT ATTAGTATTT AGCATTTGCC AGCATTTGCC AGCATTTGCC AGATACCGTC AGATACCGTC AGATACCGTC AATGACTCTA AATGACTCTA AATGACTCTA CGCAAGGCTG CGCAAGGCTG CGCAAGGCTG TTAATTTGAC TTAATTTGAC TTAATTTGAC GAGAGCTCTT GAGAGCTCTT GAGAGCTCTT TTTGTCTGGT TTTGTCTGGT TTTGTCTGGT TCGAATCATT TCGAATCATT TCGAATCATT ATAACAGGTC ATAACAGGTC ATAACAGGTC CAACGAGTTT CAACGAGTTT CAACGAGTTT GGGATAGATC GGGATAGATC GGGATAGATC AGCTTGCGTT AGCTTGCGTT AGCTTGCGTT ATGGTTTAGT ATGGTTTAGT ATGGTTTAGT 900 AATTGTCAGA AATTGTCAGA AATTGTCAGA 960 AAGGATGTTT AAGGATGTTT AAGGATGTTT 1020 GTAGTCCTAA GTAGTCCTAA GTAGTCCTAA 1080 TCAGCACCTT TCAGCACCTT TCAGCACCTT 1140 AAACTTAAAG AAACTTAAAG AAACTTAAAG 1200 TCAACACGGG TCAACACGGG TCAACACGGG 1260 TCTTGATTCT TCTTGATTCT TCTTGATTCT 1320 TAATTCCGTT TAATTCCGTT TAATTCCGTT 1380 TTAACTTCTT TTAACTTCTT TTAACTTCTT 1440 TGTGATGCCC TGTGATGCCC TGTGATGCCC 1500 ATCACCTGTA ATCACCTGTA ATCACCTGTA 1560 TTTGCAATTT TTTGCAATTT TTTGCAATTT 1620 GATTACGTCC GATTACGTCC GATTACGTCC 1680 GAGGTCTTCG GAGGTCTTCG GAGGTCTTCG Re Re Re #2 #4 #20 #2 #4 #20 1690 GATTGGCGCT GATTGGCGCT GATTGGCGCT 1750 ATTTAGAGGA ATTTAGAGGA ATTTAGAGGA TTGCAGCTGG TTGCAGCTGG TTGCAGCTGG AGTAAAAGTC AGTAAAAGTC AGTAAAAGTC CGACAGCAGT CGACAGCAGT CGACAGCAGT GTAACAAGGT GTAACAAGGT GTAACAAGGT 57 1740 GGGATGCCGA AAAGTTGATC AAACTTGATC GGGATGCCGA AAAGTTGATC AAACTTGATC GGGATGCCGA AAAGTTGATC AAACTTGATC 1790 TTCCGTTGGT GAACCTGCGG A TTCCGTTGGT GAACCTGCGG A TTCCGTIGGT GAACCTGCGG A Phylogenetic Analysis. The N81 and N88 degenerate primers amplified 1,791 base pairs of R. seeberi’s 188 SSU rDNA sequence. Of the 3 clones that were completely sequenced (Rs #2, 4, and 20) Rs# 2 was randomly chosen for the phylogenetic analysis. Phylogenetic analysis by neighborjoining and parsimony, with R. seeberi and 23 other 188 SSU rDNA sequences, supported very similar phylogenetic trees (Figure 12). In these trees, R. seeberi was always the sister taxon to the two Dermocystidium species used in this analysis. The position of the sister taxon was strongly supported with bootstrap searches in both parsimony and distance analyses. Rhinosporidium seeberi also clustered closer to other members of the DRIP clade (Ragan et al. 1996) which includes: the rosette agent, Ichthyophonus, Psorosperrnium, and both Dermocystidium sp. which were used in this study. Parsimony and neighbor analyses also showed that the DRIP group and R. seeberi are localized near the Choanoflagellates and between the kingdoms Animalia and Fungi (Figure 12). 58 Figure 12: Phylogenetic analysis by neighbor joining and parsimony, with R. seeberi and 23 other 188 SSU rDNA sequences, supported very similar phylogenetic trees. In these trees, R. seeberi was always the sister taxon to the two Dermocystidium species used in this analysis. The position of the sister taxon was strongly supported with bootstrap searches in both parsimony and distance analyses. Rhinosporidium seeberi also clustered closer to other members of the DRIP clade (Ragan et al. 1996) which includes: the rosette agent, Ichthyophonus, Psorosperrnium, and both Dermocystidium sp. which were used in this study. Parsimony and neighbor analyses also showed that the DRIP group and R. seeberi are localized near the choanoflagellates and between the kingdoms Animalia and Fungi. 59 100 a. Parsimony 68 99 60 56 Acanthocoepsis unguilata 100‘ 97 98 Dermocystidium sp. ”—T 98 Diaphanoeca grandis Dermocystidium salmonis Rhinosporidium seeberi 53 rosette agent 99 Ichthyophonus hoferi Psorospermium haeckelii Microciona prolifera Mnemiopsis Ieidyi 61 Blastocladiella emersonii Chytridium confervae 59 Glomus intraradices 51 60 Pneumocystis carinii 67 Saccharomyces cerevisiae 1‘ L' L v..., C... pombe 100 Chlorella Iobophora Chlamydomonas reinhardtii Graciiaria Iemaneiformis 100 Perkinsus marInus Perkinsus sp. Prorocentrum micans Achlya bisexualis Ochromonas danica 100 100 100 l__I b. Neighbor Joining 60 100 ‘ 100 0. 039 ..__.._.__l L__ll____ll_ll_ll choanoflagellates Mesomycetozoa animals fungi green plants red algae alveolates stramenopiles Pa SU Chapter 4 DISCUSSION Earlier studies, on R. seeben’s in vivo morphological features did not establish a definitive taxonomic relationship for this unique pathogen (Ashworth 1923, Bader and Grueber 1970, Vanbreuseghem 1973, Kannan-Kutty and Teh 1974, and Savino and Margo1983). Based on their morphological observations, these investigators suggested several possible taxonomic relationships for R. seeberi. For instance, during the last 100 years R. seeberi has been Classified as a fungi and/or as a protozoan. Rhinosporidium seeberi has also been related to other spherical protoctistans. To clarify this pathogens ambiguous phylogenetic affinities, several investigators have suggested to study R. seeberi at the molecular level. Our study indicated that the characterization of the 18S serNA of R. seeberi (1791 nucleotides), and its subsequent pyhlogenetic analysis, placed R. seeberi within the DRIP clade (DRIP: Dermocystidium, rosette agent, Ichthyophonus, and PsorospermiumXRagan et al. 1996). This study has resolved a century of taxonomic uncertainty for this human and animal pathogen. Prior to this study, all attempts to isolate nucleic acids from R. seeberi suitable for molecular studies were unsuccessful. One of the reasons for 61 previous failures may be due to the rapid degradation of R. seeberi’s nucleic acids by powerful endonucleases during cell disruption, and prior to DNA extraction, as shown in this study. The degradation process begins upon excision of the rhinosporidial polyp from the host tissue. If the sample is not processed immediately after collection, the host tissue begins to break down and release endonucleases. Our findings indicate that even when the samples were processed quickly after surgical excision, the extracted nucleic acids were invariably highly degraded. Interestingly, six faint bands were observed in our nucleic acid sample prior to RNase treatment. Some of these bands may represent R. seeberi’s rRNA precursors (458, 41 S, 328, and 208) and the fully post transcriptionally processed 188 and 288 rRNAs. The use of 188 SSU rDNA sequences to determine the evolutionary relationship between eukaryotic organisms with incomplete life cycles and tenuous morphological affinities has been very successful. In this study, we used the 18S rDNA because the frequencies of compositional changes at different positions allows the molecule to be used for comparisons over a wide spectrum of phylogenetic distances. In addition, the abundance of other 188 SSU rDNA sequences in databases enabled a more comprehensive analysis. The utility of the SSU rDNA molecule in the phylogenetic analysis of eukaryotic and prokaryotic organisms has been reviewed by Olsen and Woese 1993. Analysis of the complete sequences (3) and partially sequenced clones (19 sequences)(N85-N88deg.) of R. seeberi’s 18S rDNA, revealed that at least 9 unique copies of the 188 rDNA gene were present. This finding was surprising 62 because all clones sequenced were from the same individual. Single point mutations at 10 polymorphic sites within the 1,791 base pair amplicon were responsible for the variation observed. Due to the fact that only 19 clones were screened (3 clones completely sequenced) and 9 unique sequences have already been found, it is likely that a very diverse population of 18S SSU rDNA sequences from R. seeberi is present within the same host. One possible explanation for the variation in the 18S sequences, may be that the host was initially infected by multiple strains of the same organism, or upon repeated exposure to the source of infection (ex. the same bathing pool) became re- infected with different strains. Another possible, but unlikely event, could be that R. seeberi may be undergoing sexual reproduction during its parasitic life cycle within the infected host tissue. The results of the phylogenetic analysis indicated that R. seeberi is a member of the DRIP clade (Ragan et al. 1996), renamed MesOmycetozoa in this study. This clade is comprised of previously unknown related organisms at the most basal branch, between the animal-fungal divergence, and close to the choanoflagellates, the lowest branch of the kingdom Animalia (Kerk et al.1995, Spanggaard etal. 1996, Wainright etal.). Internal branches in the Mesomycetozoa (DRIP) clade showed strong bootstrap support by both parsimony and distance analyses, but the basal branch was strongly supported only in distance analysis. In an eariier study by Ragan et al. it was found that the DRIP’S group diverged near the animal-fungal dichotomy, although the precise position could not be conclusively resolved (Ragan et al. 1996). There 63 are two hypotheses for the position of the DRIP’s SSU rDNA’s within the eukaryote tree. Hypothesis A shows the DRIP’s having diverged after the animal-fungal dichotomy, as the most basal branch within the animal lineage. Hypothesis B shows the DRIP’s having diverged immediately basal to the animal-fungal dichotomy. Hypothesis A is the most likely and most parsimonious topology, but hypothesis B cannot be mled out statistically figure 13 (Ragan et al. 1996). Phylogenetic analyses of protein sequences and comparison of intron positions in protein-coding genes should help to determine which of these competing hypothesis is correct. OTHER ANIflALS chytr ids 1 ,‘N -r Microcionag,t wé 4‘ OTHER Choano f 1 age 1 1 ates hwy-'33TT-L féjf'éffiéffl FUNGI 0 TH ER TELI‘Q‘; Jig-:2}. ANIMALS Ch trids 2;? y A...” DRIP ’s Microciona {A OTHER I Aapusomonas Choanoflagellates FUNGI I151 A 'Sacanthamoebae 9 DRIP 8 I3 apusomonas " acanthamoebae A Figure 13: The diagrams above illustrate alternative relationships among the DRIP’s clade and other eukaryotic lineages not rejectable under the Templeton-Felsenstein and Kishino-Husegawa tests. Hypothesis A is the most likely and most parsimonious of the two. Hypothesis B, although less likely and less parsimonious, cannot be statistically ruled out (Ragan et al. 1996). This figure was adapted from Ragan et al. 1996 65 The use of molecular tools to determine the taxonomic position of an organism is of great utility and value, but the molecular data cannot stand alone. There must also be other phylogenetic characters to‘support the molecular data. In our case, it was found that members of the Mesomycetozoa (DRIP) group have several features in common with R. seeberi. For example, (i) all possess spherical parasitic stages (some have endospore-like structures) with fungus-like characteristics, which explains in part their long history of inclusion within the fungi (Elston et al. 1986). (ii) With the exception of Ichthyophonus hofen' (Spanggaard et al. 1995) they are intractable to laboratory culture; therefore, their complex life histories are unknown (Cervinka et al. 1974, Harrel et al. 1986, Hedrick et al. 1989). (iii) All are parasites related to aquatic habitats (Ajello 1998, Arsecularatne and Ajello 1998, Broz et al. 1951, Cervinka et al. 1974, Elston et al. 1986, Laveran et al. 1910, Spanggaard et al. 1995). All of these are biological and morphological characteristics that further support our phylogenetic analysis. Nucleated microorganisms and their descendants, exclusive of fungi, animals, and plants, evolved by integration of former microbial symbionts. All protoctists evolved from symbioses between at least two different kinds of bacteria (Margulis and Schwartz 1998). Modern day protoctist lineages are grouped according to their organelle structure (former symbionts). In the mitochondrion, for example, the most essential (and therefore Slowly evolving) the cristae structure can vary based upon on which symbiont it has evolved from. 66 Theses structures may be flat (as in the stramenopiles, Chytrids, and zoomastigotes), tubular (as in the alveolates), discoid (as in the amebas, slime molds, and discomitochondriates), or altogether absent (as in the archaeprotists and microsporans). The structure of the protoctistan mitochondrial cristae can provide important morphological data that can support or undo a molecular phylogenetic study. In our case, the morphological similarities between the flat cristae of the mitochondria of R. seeberi and Dermocystidium spp. supports our phylogenetic analysis. With the exception of Ichthyophonus hofen’s tubular mitochondria, all of the members of the Mesomycetozoa (DRIP’S) possessed flat mitochondrial cristae. The tubular mitochondria of I. hoferI' suggests that this organism may only be a provisional member of the Mesomycetozoa (DRIP’s) and may be more closely related to organisms that are yet to be characterized. Interestingly, the mitochondria of R. seeberi were only observed in intermediate sporangia. This may be due to the dramatic increase in cellular activity that occurs just prior to endospore formation within intermediate sporangia. They were approximately 500pm in diameter and were analogous to those of Dermocystidium species, reported by Ragan et al. 1996. Our analysis indicated that the two Dermocystidium species used in the study are a sister taxon of R. seeberi (figure 12). This molecular analogy is striking since their common morphological features were first noted by Carini in 1940 (Carini 1940). In that study, Carini suggested that the morphological features of a spherical frog parasite had similarities with R. seeberi. Because of its unusual host, however, he believed that it was different from R. seeberi and . 67 created the new genus and species Dennospon'dium hylarum (erroneously referred to as Dennospon’dium hylae in later studies). Based on Carini’s morphological description, however, we believe that the D. hylarum and Derrnosporidium granulosum infections described later (Broz et al. 1951 ) were more likely caused by R. seeberi in frogs. We based our assumption on the fact that Dermocystidium spp. and R. seeberi share phenotypic features but their habitats are different. For instance, R. seeberi has been associated with wet terrestrial habitats, while Dermocystidium spp. are found on fish in both fresh and marine waters. Although we have not examined the 18S SSU rDNA from Dermospon'dium spp., or Dermocystidium spp. growing on amphibians, we speculate that they would be better classified in the genus Rhinosporidium, or perhaps even as R. seeberi, based on their described morphological features and their capacity to cause infections on terrestrial animals. The use the 188 SSU rDNA molecule for phylogeneticanalyses has provided data to suggest the demise of an entire phylum (Siddall et al. 1995, Smothers et al. 1994) as well as to support the creation of new clades (Ragan et al. 1996). In some cases, new relationships have led to such names as the DRIP clade, an acronym derived from the names of the organisms that comprise the group. The inclusion of new members in this clade, however, renders this acronym inappropriate. Based on previous phylogenetic analyses (Ragan et al. 1996, Spanggaard et al. 1996) and the data presented in this study, we are proposing the term Mesomycetozoa (between animals and fungi) to 68 accommodate the DRIP group, R. seeberi, and any future described organisms with identical phylogenetic characteristics. This study on R. seeberi was the first to characterize, using molecular biology, a specific sequence of its genome. Additional molecular studies of this enigmatic organism must be carried to confirm this finding. The characterization of the 188 SSU rDNA of R. seeberi from humans in different geographic locations, as well as from different animals would be of great interest. The phylogenetic analyses of protein sequences and comparison of intron positions in protein-coding genes should also be done to confirm the 18S phylogeny of R. seeberi. In addition, utilizing the 188 sequence data from this study, a molecular probe specific for R. seeberi could be used to search for it’s ecological niche in nature. 69 70 APPENDIX A PHYLOGENETIC ANALYSIS OF RHINOSPORIDIUM SEEBERI’S 18S SSU RDNA GROUPED THIS PATHOGEN AMONG MEMBERS OF THE PROTOCTISTAN MESOMYCETOZOA (‘DRIP’) CLADE Roger A. Herr‘, Libero Ajelloz, John W. Taylora, Sarath N. Arseculeratne‘, and Leonel Mendoza“ Medical Technology Program, Department of Microbiology, 322 N Kedzie Lab., Michigan State University, East Lansing, MI 48824-10311. Emory University School of Medicine, Department of Ophthalmology, 1327 Clifton Road, NE, Atlanta, Georgia 30322, USA2. Department of Plant and Microbial Biology, 111 KoShland Hall, University of California, Berkeley, CA 194720-3102 USA3. Department of Microbiology, Faculty of Medicine, University of Peradeniya, Peradeniya 20400, Sri-Lanka‘. Running title: Rhinosporidium seeberi is a member of the Mesomycetozoa 71 ABSTRACT For the past 100 years the phylogenetic affinities of Rhinosporidium seeberi have been controversial. Based on its morphological features it has been classified as a protozoan or as a member of the Kingdom Fungi. We have amplified and sequenced nearly a full-length 188 (small subunit) ssu rDNA sequence from R. seeberi. Using phylogenetic analysis, by parsimony and distance methods of R. seeben’s 188 ssu rDNA and other eukaryotes, we found that this enigmatic pathogen of humans and animals clusters with a novel group of fish parasites referred to as the 'DRIP' clade (Dermocystidium, rossete agent, Ichthyophonus, and Psorospermium), near the animal-fungal divergence. Our phylogenetic analyses also indicate that R. seeberi is the sister taxon of the two Dermocystidium species used in this study. This molecular affinity is remarkable since members of the genus Dermocystidium also form spherical structures in infected hosts, produce endospores, have not been cultured, and possess mitochondria with flat cristae. With the addition of R. seeberi to this clade the acronym DRIP? is no longer appropriate. We propose to name this monophyletic clade the Mesomycetozoa to reflect the group's phylogenetic association within the Eukarya. 72 Rhinosporidium seeberi is the hydrophilic agent of rhinosporidiosis (3). This granulomatous disease of humans and animals is characterized by the development of polyps that primarily affect the nostrils and the ocular conjunctiva of its hosts. Diagnosis is essentially based on the histological detection in tissues of R. seeberr’s pathognomonic endosporulating sporangia in various stages of development. These sporangia, which are the only known phenotypic structures produced by this pathogen, range from 60-450 um or more in diameter. The mature sporangia have been estimated to contain up to 12,000 endospores (7- 15 um D) that are discharged through a pore. The liberated endospores lodge in the host's tissue and mature into endosporulating sporangia repeating its in vivo life cycle. Eariy claims that R. seeberi has been isolated in culture (4, 20) were never confirmed. The infections caused by this organism, therefore, have not been experimentally reproduced in animals and its etiologic agent has yet to be cultured (4). The taxonomy of R. seeberi has always been controversial. Seeber, who first described rhinosporidiosis in 1900, considered the sporangia of this enigmatic organism to be a sporozoan allied to the coccidia (23). Ashworth (5) in his monograph on rhinosporidiosis concluded that the nearest relatives of Rhinosporidium are not the Sporozoa but the lower fungi (Phycomycetes) such as the Chytridineae in which, sub-order, near the Olpidiaceae, Rhinosporidium is provisionally placed. Dodge (9) interpreted the sporangia of R. seeben' to be multispored asci and classified it as an ascomycetous fungus. Interestingly, early investigators also noticed the similarities that R. seeben' has with such aquatic parasites as species of Ichthyophonus (19) and Dermocystidium (7, 10), both fish pathogens that did not have a clear taxonomic background at that time. Since then, numerous investigators have incorrectly considered R. seeberi to be 73 a protozoan, a fungus and more recently a cyanobacterium (1) and a carbohydrate waste product (2). For 100 years R. seeberi has been the center of taxonomic controversies. This unsettling confusion has stemmed in part from the frustrating fact that this human and animal pathogen is intractable to culture. Thus, its life history and phylogenetic affinities have remained unknown. We are reporting in this study that phylogenetic analysis, using the 188 (small subunit) ssu rDNA of R. seeberi and 23 other microorganisms, placed the etiologic agent of rhinosporidiosis within a recently described group of fish parasites known as the DRIP clade, which we propose to rename as the Mesomycetozoa. MATERIAL AND METHODS Collection of tissue with Rhinosporidium seeberi's sporangia and endospores. Biopsed tissue, containing R. seeben’s sporangia and endospores, were obtained from two Sri Lankan men with rhinosporidiosis. The tissues were aseptically collected and transported without fixatives to the laboratory. The sporangia and endospores from both cases were processed by two different methods. In the first method the sporangia and endospores were physically dissected from the tissues, the sporangia and endospores further purified by centrifugation to remove human cells (~50 R. seeberi cells/LII). The sporangia and endospores were then disrupted with glass beads. The second method involved the use of 100 mg of human tissue containing R. seeben’s sporangia. The tissues were placed in a mortar and grinded under liquid nitrogen. Genomic DNA isolation, PCR amplification, and sequencing of Rhinosporidium seeberi's 188 ss-rDNA. In either of the above cases, the samples were placed in two eppendorf tubes and treated with SDS and proteinase K and then extracted with phenol and chloroform. The amplification of 74 the 188 ssu rDNA gene was with the polymerase chain reaction (PCR) using first the oligonucletides primers N81 (13) and from the same study the reverse primer N88. The PCR conditions were as per Gargas and DcPricst (13). Since no PCR amplicons were obtained with this set of primers, the N88 primer was degenerated as per lssakainen et al., (17): 5'TCCGCAGGTT- CACC(TA)ACGGA3'. The amplicons were subcloned into pCR 2.1-TOPO plasmids (lnvitrogen, Carlsbad, CA) purified and then sequenced using BigDye Terminator chemistry in an ABI Prism 310 Genetic analyzer apparatus (Perkin- Elmer, Foster City, CA). The sequence has been submitted to GenBank under the accession number AF118851. Before DNA extraction the sterility of the samples was always investigated by culture. Transmission electron microscopy (TEM) studies. Both of the infected human tissues, containing the sporangia and endospores of R. seeberi, were fixed in 2.5% glutaraldehyde with 0.05M Na-Cacodylate buffered saline (pH 7.4), at room temperature for 2 hours with gentle agitation. The primary fixation was followed by three 0.05M Na-Cacodylate buffered saline washes for 20 minutes each. The samples were then placed into a 1% 0804 and water solution at room temperature for 4 hours with mild agitation. 0504 post fixation was followed by three 20 minute distilled water washes and dehydration in acetone. The samples were transferred to 33% followed by 66% spurtol in acetone solutions for 30 minutes in each concentration. The samples were transferred to 100% spurtol for 5 hours, and then overnight with resin changes at the end of each time period. Ultra-thin sections were made using an ultra-microtome and sectioning the samples into 100 nm sections. The grids with the ultra thin sections were post stained using uranyl acetate for 30 minutes followed by Hanaichi et al., lead (14) for 3 minutes. After the post staining procedure a thin layer of carbon was 75 evaporated onto the surface of the grids. Examination of ultra-thin sectioned material was with a Phillips CM-10 electron microscope. Phylogenetic analysis. The 188 ssu rDNA sequence of R. seeberi was aligned by eye with sequences previously analyzed by Spanggaard et al. (27) and Ragan et al. (22) using Sequence Navigator (Perkin Elmer/Applied Biosystems). Regions of ambiguous alignment were excluded from phylogenetic analysis, and gaps introduced to facilitate alignment were treated as missing data. Phylogenetic analysis was by a distance method (neighbor joining using Kimura's 3 parameter multiple hit correction) (27) and parsimony. Support for internal branches was assessed using 1000 bootstrapped data sets. Parsimony analyses were conducted with the computer software program PAUP (PAUP: Phylogenetic analysis Using Parsimony, 3.1; D.L. Swofford, Illinois Natural History Survey, Champlain, IL). Parsimony analysis employed 1000 heuristic searches with randomized orders of taxa entry to increase the chance of finding the shortest trees. Trees were drawn with stramenopiles and alveolates as the outgroups, based in previous phylogenetic studies (18, 22, 27). RESULTS Phylogenetic analysis using Rhinosporidium seeberi’s 188 ssu rDNA. The N81 and the reverse modified primer amplified 1790 bp of the gene from both cases, nearly a full length of the R. seeben’s 188 ssu rDNA sequence. Sequence analysis of R. seeben’s 1790 bp amplicons showed that the DNA sequences from both rhinosporidiosis cases were identical. Phylogenetic analysis by neighbor joining and parsimony, using R. seeberi and 23 other 188 ssu rDNA sequences, supported very similar phylogenetic trees (Fig. 1). In these trees, R. seeberi was always the sister taxon to the two Dermocystidium species used in this analysis. The position of the sister taxon was strongly supported with bootstrap search in both parsimony and distance analysis. 76 Rhinosporidium seeberi also clustered closer to other members of the 'DRIP’ clade, the rosette agent, Ichthyophonus, and Psorospermium, used in this study. Parsimony and neighbor analyses also showed that the DRIP group and R. seeberi are localized near the choanoflagellates hand between the kingdoms Animalia and Fungi (Fig. 1). TEM analysis. Transmission electron microscopy analysis from both tissues showed the typical sporangial phenotype that characterized R. seeben‘s infections. Sporangia at different stages of development were observed throughout the infected tissues. Rhinosporidium seeben’s mitochondria were difficult to find in any of the phenotypic stages except the intermediate sporangia. The intermediate sporangial stage was characterized by the presence of a prominent cell wall, multiple nuclei, numerous mitochondria, and laminated bodies (Figure 2 A). Our TEM analysis showed that R. seeberi has mitochondria with flat cristae (Fig. 2 B). DISCUSSION Our phylogenetic analysis indicated that R. seeberi is a member of the 'DRIP’ clade (22), renamed Mesomycetozoa in this study. This clade comprises previously unknown related organisms at the most basal branch between the animal-fungal divergence and close to the choanoflagellates the lowest branch of the kingdom Animalia (18, 27, 28). lntemal branches in the Mesomycetozoa (DRIP) clade showed strong bootstrap support by both parsimony and distance analyses, but the basal branch was strongly supported only in distance analysis. Interestingly, members in the Mesomycetozoa (DRIP) group have several features in common with R. seeberi. For instance, a) all posses spherical parasitic stages (some have endospore-like structures) with fungal-like characteristics, which explain in part their long history of inclusion within the fungi (12), b) with the exception of Ichthyophonus hoferi (26), they are intractable to 77 laboratory culture, therefore their complete life histories are unknown (8, 15, 16), and C) all are parasites related to aquatic habitats (3, 4, 6, 8, 12, 19, 26), all biological and morphological characteristics that further support our phylogenetic analysis. In addition, the morphological similarities of R. seeben’s and Dermocystidium spp’s mitochondria with flat cristae also support our phylogenetic analysis. Our analysis indicated that the two Dermocystidium species used in our study are a sister taxon of R. seeberi (Figure 1). This molecular analogy is striking since their common morphological features were first noted by Carini in 1940 (7). In that study Carini suggested that the morphological features of a spherical frog parasite had similarities with R. seeberi. Because of its unusual host, however, he believed that it was different from R. seeberi and created the new genus and species Dermospon‘dium hylarum (erroneously referred to as D. hylae in later studies). Based on Carini's morphological description, however, we believe that D. hylarum and D. granulosum, described later (6), were more likely cases caused by R. seeberi in frogs. We based our assumption On the fact that both Dermocystidium spp and R. seeberi share phenotypic features, but their habitats are different. For instance, R. seeberi has been found in wet terrestrial habitats, while Dermocystidium spp are found on fish in both fresh and marine waters. Although we have not examined the 188 ssu rDNA sequence of Dennospon’dium spp or Dermocystidium spp growing on amphibians, we speculate that they would be better classified in the genus Rhinosporidium, perhaps even as R. seeberi, based on their described morphological features and their capacity to cause infection on terrestrial animals. The use of 188 ssu rDNA sequences to determine the evolutionary relatedness between eukaryotic organisms with incomplete life cycles and tenuous morphological affinities has been very successful. The usefulness of 78 the ssu rDNA molecule in the phylogenetic analysis of eukaryotic and prokaryotic organisms has been reviewed by Olsen and Woese (21). In our study we used the 188 ss rDNA because the frequencies of compositional changes at different positions allow the molecule to be used for compariSons over a wide spectrum of phylogenetic distances. In addition, the abundance of other 188 ssu rDNA sequences in databases enables a better comprehensive analysis. Moreover, phylogenetic analysis using this molecule has provided data to suggest the demise of an entire phylum (24, 25) or to support the creation of new clades (22). In some cases new relationships have lead to such names as the DRIP? clade, an acronym derived from the organisms that comprise the group. The inclusion of new members in this clade, however, renders this acronym inappropriate. Based on previous phylogenetic analyses (22, 27) and the data presented in this study, we are proposing the term Mesomycetozoa (between animals and fungi) to accommodate the ‘DRIP’ group, R. seeberi, and any future described organisms with identical phylogenetic characteristics. The finding that R. seeben’s 188 ssu rDNA sequences amplified from tissues of both cases were identical suggested that Rhinosporidium is a monotypic genus. However, new 188 ssu rDNA sequences from other humans and animals with rhinosporidiosis have to be evaluated to validate our study. It is anticipated that the inclusion of a human and animal pathogen within the Mesomycetozoa will encourage more studies of this clade. It is our hope that the results of such studies will unveil more details about the ecology, biology, and molecular aspects of this unique group of microorganisms, recently suggested as the possible ancestors from which the fungi and the animals may well have originated. 79 ACKNOWLEDGMENTS We thank Kathy Lee for her involvement in the evaluation of the degenerated reverse primer on the stramenopilan Pythium insidiosum that eventually led to its use on R. seeben’s DNA. We also thank John Gerlach for his help and suggestions during these analyses as well as John Heckman and Sally Burns at the Center for Electron Optics, MSU, for their assistance. This study was supported in part by the Medical Technology Program, MSU. 80 REFERENCES 1. Ahluwalia, K. B., N. Maheshwari, and R. C. Deka. 1997. Rhinosporidiosis: A study that resolves etiologic controversies. Amer. J. Rhinol. 11:479-483. 2. Ahluwalia, K. B. 1992. New interpretations in rhinosporidiosis, enigmatic disease of the last nine decades. J. Submicrosc. Cytol. Pathol. 24:109-114. 3. Ajello, L. 1998. Ecology and epidemiology of hydrophilic infectious fungi and parafungi of medical mycologycal importance: A new category of pathogens, p. 67-73, Vol. 4, chap. 4. In L. Ajello, th.R J. Hay, (Eds), Topley & Wilson?s Microbiology and Microbial Infections, 9th ed. Arnold, London. 4. Arsecularatne, S. N., and L. Ajello. 1998. Rhinosporidium seeberi. p. 596- 615, Vol.4 Chap. 31. In L. Ajello, R.J. Hay, (Eds.,) Topley & Wilson?s Microbiology and Microbial Infections, 9th ed. Arnold, London. 5. Ashworth, J. H. 1923. On Rhinosporidium seeberi (Wemicke, 1903) with Special reference to its sponIlation and affinities. Trans. Roy. Soc. Edin. 53:301-342. 6. Broz, O. and M. Privora. 1951. Two Skin parasites of Rana temporan'a: Dermocystidium ranae Guyenot and Naville, and Dermospon’dium granulosum n.sp. Parasitology 42:65-69. 7. Carini, A. 1940. Sobre um parasito semlhante ao ‘Rhinosporidium’, encontrado em quistos da pele de uma ‘hyla’. Arquivos lnstituto Biologico, Sao Paulo, Brazil, 11:93-96. 8. Cervinka, 8., J. Vitovec, J. Lom, J. Hoska, and F. Kubu. 1974. Dermocystidiosis: A gill disease of the carp due to Dermocystidium cypn'ni n.sp. J. Fish Biol. 6:689-699. 9. Dodge, C. W. 1935. Rhinosporidium. In Medical Mycology. Fungous Diseases of Men and other Mammals. p. 151-152. CV. Mosby Co., St. Louis. 10. Dunkerly, J. S. 1914. Dermocystidium pusula Perez, parasitic on trutta fario. Zool. Anz. 44:179-182. 11. Dykova, l and J. Lom. 1992. New evidence of fungal nature of Dermocystidium koi 1 950. J. Appl. Ichthyol. 8: 1 80-1 85. 81 12. Elston, R. A., L. Harrell, and M. T. Wilkinson. 1986. Isolation and in vitro characteristics of Chinook salmon (Oncorhynchus tshawytscha) rosette agent. Aquaculture 5621-21. 13. Gargas, A. and P. T. DcPricst. 1996. A nomenclature for fungal PCR primers with examples from intron-containing SSU rDNA. Mycologia 88:745- 748. 14. Hanaichi,T., T. Sato, T. lwamoto, J. Malavasi-Yamashiro, M. Hoshlno, and N. Mlzuno, N. 1986. A stable lead by modification of Sato's method. J. Electron. Microsc. 35:304-306. 15. Harrell, L. W., R. A. Elston, T. M. Scott, and M. J. Wilkinson. 1986. A significant new systemic disease of Net-Pen reared Chinook salmon (Oncorhynchus tshawytscha) brood stock. Aquaculture 55:249-262. 16. Hedrick, R. P., C. S. Friedman, and J. Modin. 1989. Systemic infections in atlantic salmon Salmo salar with a Dennocystidium-like species. Dis. Aquat. Org. 7:171-177 17.Issakainen, J., J. Jalava, E. Eerola, and C. K. Campbell. 1997. Related ness of Pseudallescheria, Scedospon'um, and Graphium pro parte based on SSU rDNA sequences. J. Med. Vet. Mycol 35:389-398. 18. Kerk, D., A. Gee, M. Standish, P. O. Wainwright,_A. 8. Drum, R. A. Elston, and M. L. Sogin. 1995. The rosette agent of Chinook salmon (Oncorhynchus tshawytscha) is close related to choanoflagellates, as determined by the phylogenetic analysis of its small ribosomal subunit RNA. Marine Biol. 122:187-192. 19. Laveran, A. and A. Pettlt. 1910. Sur une epizootie des truites. CR. Acad. Sci. Paris 151:421-423. 20. Levy, M.G., D. J. Meutem, and E. B. Breitschwerdt. 1986. Cultivation of Rhinosporidium seeberi in vitro: Interaction with epithelial cells. Science 234:474-476. 21.0lsen, G.J and CR. Woese. 1993. Ribosomal RNA: a key to phylogeny. FASEB Journal. 7:113-123. 22. Ragan, M. A., C. L. Goggin, R. J. Cawthom, L. Cerenius, A. V. C. Jamieson, S. M. Plourdes, T. G. Tand, K. Soderhall, and R. R. Gutell. 1996. A novel clade of protistan parasites near the animal-fungal divergence. Proc. Natl. Acad. Sci. 93:11907-11912. 82 23. Seeber, G. R. 1900. Un nuevo esporozuario parasito del hombre. Dos casos encontrados en polypos nasales. Thesis, Imprenta Libreria "Boullosa", Buenos Aires, Argentina. 24. Siddall, M. E., D. S. Martin, D. Bridge, S. S. Desser, and D. K. Cone. 1995. The demise of a phylum of protists: Phylogeny of myxozoa and other parasitic cnidaria. J. Parasitol. 81:961—967. 25. Smothers, J. F., C. D. von Dholen, L. H. Smith Jr., R. D. Spall. 1994. Molecular evidence that the myxozoan protists are metazoans. Science 265:1719- 1721. 26. Spanggaard, B., H. H. Huss, and J. Bresciani. 1995. Morphology of Ichthyophonus hoferi assessed by light and scanning electron microscopy. J. Fish Dis. 18:567-577. 27. Spanggaard, B., P. Skounoe, L. Rossen, and J. W. Taylor. 1996. Pylogenetic relationships of the intracellular fish pathogens Ichthyophonus hoferi, fungi, choanoflagellates, and the rosette agent. Marine. Biol. 26:109-115. 28. Wainright, P. 0., G. Hinkle, M. L. Sogin, and S. K. Stickel. 1993. Monophyletic origins of the metazoa: An evolutionary link with fungi. Science 260:340-342. 83 APPENDIX B LOCALIZATION OF AN IMMUNOREACTIVE INNERWALL ANTIGEN OF RHINOSPORIDIUM SEEBERI EXPRESSED ONLY DURING MATURE SPORANGIAL DEVELOPMENT R.A. HERR,1 L. MENDOZA," S.N. ARSECULERATNE,2 AND L. AJELLO° Michigan State University, Medical Technology Program, East Lansing, MI,1 Faculty of Medicine University of Peradeniya, Sri-Lanka,2 Emory Eye Center, Emory University, Atlanta, GA3. Running title: An lmmunoreactive Antigen of Rhinosporidium seeberi Rhinosporidium seeberi infections are characterized by the development of polyps primarily localized in the mucous membranes of its hosts. Although numerous light and electron microscopic studies have been performed in the past 40 years, nothing is known of the antigens involved in its infections. We investigated the localization of R. seeberi antigens during infection using serum and infected tissue from a Sri Lankan patient with rhinosporidiosis. The tissue was fixed in white resin, thin sectioned, fixed onto nickel grids and evaluated by EM for the presence of spherules in different stages of development. The tissues were reacted with the patient’s serum and then labeled with 10 nm of protein A colloidal gold. After labeling, the tissues were post-stained with uranyl acetate and KMnO4. It was found that the gold particles fixed to antibodies that recognized the second inner mucoid-like cell wall layer of mature sporangia. Minor gold binding was also detected on the first inner cell wall layer in close contact with the cell wall and to the capsule of immature and mature endospores. Strikingly, the cytoplasm and other structures of the endospores before and after their release did not undergo gold labeling. Our results suggest that this inner wall mucoid-like antigen is only expressed in mature spherules during the process of endospore cleavage or in spherules with well-developed endospores. The hosts come in contact with this mucoid-like substance after release. This may explain why immature endospores are not labeled by the gold particles. This is the first time that an antigen with a potential role in the immunology of R. seeberi infections has been detected. 85 INTRODUCTION Rhinosporidiosis is a chronic disease of humans and other animals caused by Rhinosporidium seeberi, The disease is characterized by the formation of polyps on the mucous membranes of the infected host. Rhinosporidiosis is not a life threatening disease and its treatment basically is limited to surgical removal of polyps. Relapses, however, are common. Rhinosporidiosis is most prevalent in Asia, but the disease has been reported in all other continents except Australia. Attempts to isolate R. seeberi from infected tissues have not yet succeeded, thus the phylogenetic classification of this organism -fungus or protozoan- is unclear and R. seeberi=s ecologic niche and life cycle in nature remain unknown. Recently, we conducted immunolectron microscopic studies to investigate the in vivo development of R. seeberi and the immunology of its parasitic state. The results of our studies have shown that serum from an individual with rhinosporidiosis contains antibodies against an antigen present only in the mucoid-like inner wall of R. seeberis mature sporangia. Our electron microscopic study also indicated the presence of activated macrophages engulfing or attempting to engulf different stages of R seeberi. These findings affimi that R. seeberi is an organism that has evolved to parasitize its hosts and that the hosts develop antibodies against specific epitopes present in the mature sporangia of this organism. 86 MATERIALS AND METHODS Tissue and serum samples. Clinical material for this study was obtained from a severe case of rhinosporidiosis in a Sri- Lankan human patient. Portions of the infected tissues were fixed in glutaraldehyde and formaldehyde and a serum sample preserved with merthiolate (0.02%). Fixation - Dehydration - Embedding of Tissue for EM studies. Previous to immunolabeling the tissue samples undenNent fixation, dehydration, and embedding. Briefly, the tissue samples were removed from the primary fixative and washed with TTBS (1% w/v bovine serum albumin, 0.02% sodium azide, and 0.05% Tween 20 in Tris-buffered saline) for 10-20 minutes (3X) at 25°C. The tiSsue samples were then transferred to 10% ethanol agitated for 10 minutes, then transferred to 20% ethanol and agitated for 10 more minutes. The washed tissue was dehydrated further in 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 100% ethanol each step for 10 minutes with slight agitation. The tissue samples were transferred to fresh 100% ethanol and held for an hour. The dehydrated samples were infiltrated with 25% LR White (medium) resin for 30 minutes with agitation, and then transferred to 50% LR White resin and agitated for 30 more minutes. Infiltration of the tissue samples continued with 75%, 100%, and another 100% LR White resin concentrations for 2 hours, 2 hours, and 12 hours respectively. The tissue was then cast in fresh resin at 65?C for 48 hours. 87 Immunolabeling of tissue samples. Embedded tissue was ultra-microtomed (100-130nm) onto nickel grids for electron microscopic examination. The tissue sections on the grids were blocked with TTBS for 15 minutes. The tissue sections were allowed to react with 50% patient serum diluted in TTBS with BSA at 25?C for an hour. After incubation, the grids holding the tissue sections were carefully rinsed in TTBS. Then, the tissue sections were incubated for 1 hour at 25°C in 10% protein A colloidal gold (10 or 25 nm in diameter) diluted in TTBS. Any unbound Protein A colloidal gold was removed by a gentle rinse with TTBS followed by a brief rinsing with water. Post staining of the immunolabeled tissue sections was carried out with a 4:1 solution of uranyl acetate: KMnO4 (1% in H20) for 5 minutes. A light carbon coat was applied to the immunolabelled tissue sections on the grids before examination. RESULTS lmmunolocalization of R. seeberi Antigens. Tissue samples prepared for immunolabeling by dehydration in ethanol and infiltration with LR White resin were thin sectioned onto nickel grids and then immunolabelled as per Materials and Methods. Tissue sections on the grids were reacted with serum from a patient with rhinosporidiosis. It was found that most of the protein A-colloidal gold particles were bound to antibodies that specifically recognized the mucoid-like second layer beneath the cell wall in mature sporangia with three mucoid-like layers (Figure 6 a, b). A few colloidal gold particles were also found on the first 88 inner wall layer, in the endospores' cytoplasm, and on the mucoid-like capsule of immature and mature endospores. Immunolabeling of the third mucoid-like fibrilar layer that is in close contact with endospores was not observed (Figure 7 a, b). In mature sporangia with only two layers beneath the cell wall, the colloidal gold particles were detected only in the layer in contact with the sporagiums' cell wall. No immunolabeling was observed on the second layer in contact with the immature endospores (Figure 8 a, b, c). This finding suggested that the first layer underwent further differentiation to become the two electron dense mucoid-like layers closer to the cell wall in later developmental stages of the mature sporangia. The juvenile and intermediate sporangia did not undergo immunolabeling. Similarly, human host tissue and negative control tissue did not undergo specific immunolabeling. DISCUSSION. The majority of the juvenile sporangia were in contact with the host’s subcutaneous tissues and just a few were found within macrophages. The cytoplasm of the phagocytized juvenile sporangia was more electron dense than that of the non-phagocytized released endospores suggesting a change that precedes sporangial destruction had occurred. Active cell mediated immune response had been previously documented in cases of humans with rhinosporidiosis (Chitravel). Mature sporangium with only one inner wall layer usually had a hard thick 89 cell wall that was difficult to out without deformation. This hard thick cell wall originated directly from the electron dense cell wall of the intermediate sporangium by differentiation into a less electron dense structure. Interestingly, gold particles were found specifically bound to this inner wall structure suggesting that the presence of an antigen in the inner wall no expressed in early stages of sporangia has been formed. The presence of this hard inner cell wall suggests that during this stage deep structural changes have occurred in the sporangia cell wall. The thick hard cell wall was found to be involved later in the formation of the two other inner mucoid-like wall layers in the following developmental stage of the mature sporangia. We base this assumption on the fact that at this stage a fine electron dense wall-like structure is evident in the outer most part of this structure and in that its translucent features remain the inner layers observed in the later stages. Sporangia with two inner wall layers were Characterized by the presence of a prominent translucent inner wall layer in contact with the cytoplasm and a more electron dense inner layer in contact with a thin cell wall. Gold particles were bound only to the inner layer in contact with the cell wall. The inner layer in contact with the cell wall differentiate into two inner layers, a mucoid-fibril, which remain in contact with the cell wall, and a central mucoid-Iike layer between the two fibril-like inner layers. Gold particles were heavily bound to the central inner layer. less strongly to the inner layer in contact to the cell wall. No gold particles were found on the inner wall in contact with the endospores. The results of our immunoelectron microscopic study, using serum from a 90 patient with rhinosporidiosis and Protein A colloidal gold, showed that the serum contained antibodies against a specific inner wall antigen of R. seeberi. The antigen was found expressed only in mature sporangia. The detection of minor gold biding against the capsule and other cytoplaSmic structures indicated that the patient’s serum may also contain low concentrations of antibodies against these structures. Previously published serological studies, using disrupted R. seeberi endospores as antigen in inmmunodiffusion (ID) and counterimmunoelectrophoresis (CIE) tests, failed to demonstrate the presence of circulating antibodies in 69 patients with active rhinosporidiosis. The failure to detect antibodies in that study can be attributed to the fact that the major antigen, detected by the serum used in our immune-electron microscope study, was exclusively localized within the cell wall of mature sporangia and not in the endospores. In addition to this, they used two insensitive tests ID and CIE, both inadequate to detect low concentrations of antibodies. Although, their study failed to confirm antibodies in humans with rhinosporidiosis, a circulating R. seeberi antigen was consistently present in their test sera, but it was not characterized. We did not investigate the presence of this antigen in our serum sample. This is the first time that circulating antibodies against R. seeberi have been demonstrated and first time that a specific antigen of relevance in the immunology of R. seeberi infections has been reported. In addition, it has been shown that cellular immunity plays a role in controlling infection. Macrophages engulfing released endospores lead us to believe that despite the destruction of some of R. seeben’s endospores by macrophages, the host’s immune system is 91 overwhelmed by the number of endospores released by mature sporangia, thus ensuring the parasite's survival. Recently reported studies by Ahluwalia (1991, 1992) and Ahluwalia et al., (1994) maintain that the in-vivo tissue form of R. seeberi is not a sporangium of a “fungus” but a unique nodular body organized in host tissue for the elimination of two indigestible carbohydrates: an exogenous polysaccharide and defective proteoglycans originating from a diet of tapioca. This unconventional conclusion was concurred in by Azadeh et al., (1994). They hold that the 'sporangia" and “spores” of R. seeberi are “lysosomal bodies loaded with indigestible residues to be Cleared via transepithelial elimination or segregated/destroyed by secondary immune/granulomatous responses”. The authors go on to state that “it is not clear what pathobiological demands or influences motivate assembly of waste products in such an organized and apparently unprecedented manner”. This bizarre hypothesis was conceived in the belief among others, that no organism has been isolated from infected tissue, that antibodies against this organism have not been yet detected in individuals with rhinosporidiosis, and that the available antifungals are ineffective against R. seeberi. The data presented in our study, however, indicate that patients with rhinosporidiosis have active humoral and cellular responses to R. seeberi’s sporangia. Our electron microscopic images also revealed the presence of nuclei with a prominent nucleolus in endospores as well as in sporangia. These findings confirm previous observations and sustain the belief that R. seeberi is a eukaryotic pathogen and not a self assembled carbohydrate artifact as has been postulated. 92 93 References 1. Ahluwalia, KB. 1992. Plant molecule in human disease - a novel association. In: Wegmann RJ. and Wegmann, M.A., eds. Gene regulation and molecular aspects of muscle, liver, pancreas, connective tissue plants. Peelers Press, Leuven, Belgium, pp289-292. 2. Ahluwalia, KB. 1992. New interpretations in rhinosporidiosis, enigmatic disease of the last nine decades. J. Microscopic Cytol. and Pathol. 24:109- 114. 3. Apple, D.J. 1983. “Papillome” der conjunktiva bedingt durch rhinosporidiose. Fortschr Ophthalmol. 79:571-574. 4. Azadeh, B., N. Baghoumian, and CT El-Barkri. 1994. Rhinosporidiosis: immunochemical and electron microscopic studies. J. Laryngology and Otology 108: 1 048-1 054. 5. Chitravel, V., T. Sundararaj, 8. Subramanian, M. Kumaresan, and S. Kunjithapadam. 1982. Detection of circulating antigen in patients with rhinosporidiosis. Sabouraudia 20:185-191. 6. Chitravel, V., T. Sundararaj, S. Subramanian, M.‘ Kumaresan, and S. Kunjithapadam. 1981. Cell mediated immune response in human cases of rhinosporidiosis. Sabouraudia 19:135-142. 7. Kannan-Kutty, M. and JB Gomez. 1971. The ultrastructure and life history Of Rhinosporidium seeberi. Southeast Asian J. Trop. Med. Publ. Health 229-16. 8. Kannan-Kutty, M. and EC. Teh. 1974. Rhinosporidium seeberi: an electron microscopic study of its life cycle. Pathology 6:63-70. 9. Kannan-Kutty, M. and EC. Teh. 1975. Rhinosporidiun seeberi: an ultrasctructural study of its endosporulation phase and trophocyte phase. Arch. Pathol. 99:51-54. 10. Kennedy, F.A., R.R. Buggage, and L. Ajello. 1995. Rhinosporidiosis: a description of an unprecedented outbreak in captive swans (Cygnus spp.) and a proposal for revision of the ontogenic nomenclature of Rhinosporidium seeberi. J. Med. Vet. Mycol. 33:157-165. 11. Moses, J.S. C. Balachandran and more 1991. Rhinosporidium seeberi: light, 94 phase contrast, fluorescent, and scanning electron microscopic study. Mycopathologia 1 14: 1 7-20. 12. Savino D.F. and CE. Margo. 1983. Conjuntival rhinosporidiosis light and electron microcoplc study. Ophthalmol. 90:1482-1489. 13. Teh, EC. and M. Kannan-Kutty. 1975. RhinOsporidium seeberi: spherules and their significance. Pathology 7:133-137. 14. Thianprasit, M. and K Thagemgpol. 1989. Rhinosporidiosis. Curr. Top. Med. Mycol. 3:64-85. 15. Vanbreuseghem, R1973. Ultrastructure of Rhinosporidium seeberi. Int. J. Derrnatol. 12:20-28. 95 APPENDIX C IN VITRO STUDIES ON THE MECHANISMS OF ENDOSPORE RELEASE IN RHINOSPORIDIUM SEEBERI LEONEL MENDOZA,1. ROGER A. HERR,1 SARATH N. ARSECULERATNE,2 AND LIBERO A.IELLO3 Medical Technology Program, Department of Microbiology. Michigan State University, East Lansing, Ml,1; Faculty of Medicine, University of Peradeniya, Peradeniya, Sri-Lankaz; Emory University School of Medicine , Department of Ophthalmology, Atlanta, GA3 Running title: Endospore release in Rhinosporidium seeberi 96 Studies of Rhinosporidium seeberi have demonstrated that this organism has a complex life cycle in infected tissues. Its in vivo life cycle is initiated with the release of endospores into a host’s tissues from its spherical sporangia. However, little is known about the mechanisms of sporangium formation and endospore release since this pathogen is intractable to culture. We studied the in vitro mechanisms of endospore release in R. seeberI's endospores from viable sporangia. It was found that endospore discharge begins with an increase in the lntemal pressure of the sporangia, a rearrangement of the endospores, and the formation of an apical pore located in the cell wall of R. seeben’s mature sporangia. Only one pore per sporangium is formed. Our study also found that these events were stimulated by water. The finding of early stages of pore formation in juvenile and intermediate sporangia suggested that their formation is genetically programmed and is not a random process. The stimulation of R. seeberis sporangia by water supports the epidemiological studies that had linked this pathogen with wet environments. It also explains, in part, its affinities for mucous membranes in the infected hosts. The microscopic features of endospore discharge suggest a connection with organisms classified in the Kingdom Protoctista. Our study strongly support a recent finding that placed R. seeberi with organisms in the protoctistan Mesomycetozoa clade. Rhinosporidiosis is a disease of subcutaneous tissues, particulariy the mucous membranes, caused by Rhinosporidium seeberi, a eukaryotic organism recently classified within the new protoctistan Mesomycetozoa clade (5, 12, 14). Histopathological and Ultra structural studies of R. seeberi have demonstrated that it has a complex life cycle in infected tissues (6-9, 13, 14, 17). According to those studies, the life cycle is initiated with the release of endospores in a host’s tissues from spherical sporangia (100 to 450 um D). Once implanted, the 97 endospores increase in size and progressively develop into juvenile, intermediate, and finally mature sporangia with endospores (5, 14). The endospores are then released and the in vivo cycle is reinitiated. Some authors have suggested that the release of the endospores occurs by mechanical rupture of R. seeberI‘s sporangial cell wall, as found in Coccidioides immitis (15). Based on histological studies others, however, have postulated that endospores are discharged through a pore that develops in the sporangium's cell wall (4, 7, 13, 15, 19). We studied the in vitro mechanisms of endospore release in fresh sporangia of R. seeberi and found that water stimulated several events inside the sporangia that culminated with endospore discharge. We found that the release of endospores from the mature sporangia of R. seeberi is a well organized and dynamic process. MATERIALS AND METHODS Tissue samples. Tissues samples were obtained from two Sri Lankan men with subcutaneous rhinosporidiosis. The biopsied tissue was surgically removed, immediately iced and transported to the laboratory. Small portions of the tissues (100 mg) were fixed in formaldehyde and glutaraldehyde for histopathology and ultrastructural studies respectively. Induction of endospores release from mature sporangia. The tissues were microscopically dissected, using an inverted microscope, to remove sporangia in different stages of development. The sporangia then were placed on glass slides with or without distilled water and observed for six hours or more with a light microscope. In addition, non-dissected pieces of tissue (3 mm D) were simultaneously studied. The nomenclature recently proposed by Kennedy et al. (14) was adopted to conform with their Up-dated terminology for R. seeben's in vivo developmental 98 stages. The events leading to endospore release were also video taped using a Sony video camera attached to a BH2 Olympus microscope. Light and electron microscopic studies. The tissues containing sporangia and endospores of R. seeberi were fixed in 2.5% glutaraldehyde with 0.05M Na-cacodylate buffered saline (pH 7.4), at room temperature for 2 hours with gentle agitation. The formalin fixed tissues were embedded in paraffin, sectioned, and stained with haematoxylin and eosin (H&E). The primary fixation was followed by three 0.05M Na-cacodylate buffered saline washes of 20 minutes each. The samples were then placed in a 1% 0804 and water solution and held at room temperature for 4 hours with mild agitation. OsO4 post fixation was followed by three 20 minute distilled water washes and dehydration in acetone. These samples were transferred to 33% spurtol followed by 66% spurtol in acetone solutions for 30 minutes at each concentration. The samples then were transferred to 100% spurtol for 5 hours, and held overnight with resin changes at the end of each time period. Ultra-thin sections were cut using an Ultra-microtome into 100 nm sections. The grids with the sections were post stained using uranyl acetate for 30 minutes followed by Hanaichi's lead (10) for 3 minutes. The sectioned material was examined with a Phillips CM-10 electron ‘ microscope. RESULTS Microscopic features of fixed sporangia in light and electron microscopy. Numerous sporangia at different stages of development from both cases were observed in H&E tissue sections. Transmission electron microscopic (TEM) studies of mature sporangia with endospores showed that they possessed a thin cell wall, mature endospores with a capsule, an electron lucent mucous- like substance near the exit pore, and a genninative zone opposite to the exit pore. Juvenile and intermediate sporangia were Characterized by their smaller 99 sizes, a prominent cell wall, and the presence of numerous mitochondria, nuclei and laminated bodies. Microscopic features of fresh sporangia on wet mount preparations. Fresh sporangia collected by mechanical manipulation directly from the Clinical specimens, revealed spherical sporangia approximately between 100 um and >450 um in diameter and numerous free endospores ~5 um to ~8 um in diameter. Most sporangia, especially those that were >450 um in diameter, were filled with round to oval endospores. No capsule was evident on free and/or lntemal endospores under light microscopy. The small sporangia (<100 um in diameter) were filled with a granular cytoplasmic substance but did not contain endospores. Cytoplasmic organelles were not evident under light microscopy in those sporangia. Some sporangia with a granular cytoplasmic content showed the presence of early stages of pore development. Process of endospore release. Approximately 80% of the mature sporangia with endospores were visibly stimulated and initiated endospore release following the addition of water. Immature sporangia with granular cytoplasmic contents were not stimulated. It was found that the process of endospore release started when the apical pore had fully formed. Only one pore per sporangium was formed. The ring Shaped pore was ~10.0 um in diameter and in some sporangia they were wider than ~12.0 um. The process of endospore release started with the ejection of endospores to a distance of several micrometers away from the pore. The endospores were not released steadily through the pore, but were ejected one by one or in small groups of several endospores, sometimes numbering ~80 endospores. In a few instances, particularly in the early stages of discharge, the endospores were ejected so violently that the sporangium spun around its axis. In those cases, the release of more than 1,000 endospores occurred in a lapse of only a few seconds. 100 Emission of mucoid material from the sporangium always accompanied the endospores on their way out. Immediately under the pore, a mucoid plug-like structure was invariably present. This mucoid structure seemed to regulate endospore discharge. Once endospore ejection had been initiated, the endospores below this plug-like structure were rearranged to occupy the space once held by the released endospores. This event created an lntemal movement of the endospores from the lower half of the sporangium toward the apical pore. Under our experimental conditions, the sporangia remained spherical at all times. Even in those cases where more than 10,000 endospores had been released, the sporangia maintained their spherical forms. A few minutes before endospore release, all of the spherical mature endospores were located in the upper half of the sporangium (facing the pore), whereas the less differentiated endospores, usually small (2.0 um to 5 um in diameter) and oval in form, were found in the lower half of the sporangium. The immature endospores seemed to have developed in an area near the cell wall known as the 'germinative zone”. The small, oval, immature endospores increased in size and had matured by the time they reached the center of the sporangium. This process took approximately two hours to complete. All of the sporangia during the release process, contained endospores at different stages of maturity inside the sporangia, even 24 hours after the beginning of the release process. Effect of water on sporangia. Because water seemed to stimulate the release of endospores from mature sporangia, several sporangia were picked out of the infected tissues and placed on glass slides without water. Under these conditions the endospores were not released. When a drop of sterile distilled water was added to these sporangia it was found that ~80% of the mature 101 sporangia with endospores began discharging a steady stream of endospores. In some case this event was so violent that the sporangium spun several times around its axis. Likewise, when small, 3 mm in diameter pieces of the infected tissue was placed in contact with sterile distilled Water, some of the sporangia with mature endospores begins lodging their spores inside the tissue, as previously noted. DISCUSSION When Seeber (18) first described infections caused by R. seeberi, he believed that the etiologic agent was a protozoan of the coccidia group. Later other researchers, however, based on electron microscopic, silver and periodic acid schiff (PAS) studies, postulated that R. seeberi had more features in common with fungi than the coccidia. Since then numerous investigators erroneously classified R. seeberi either as a protozoan or a fungus and even as a blue-green bacterium or a carbohydrate waste (1, 2, 6, 8, 13). A recent study based on 188 ss-rDNA, however, indicated that R. seeberi is phylogenetically associated with members of the protoctistan Mesomycetozoa clade (12). Interestingly, all of the morphological, epidemiological and biological features that R. seeberi has in common with other members of this clade are in agreement with the molecular data. As reported in this study, R. seeberi requires the presence of water for endospore release. This concept, however, is not entirely new. In London Ashworth in 1921 (6), working with fresh tissue samples obtained from a resident Indian student with rhinosporidiosis, found that water and a variety of watery substances (mucous secretions, ringer solution, etc.) stimulated the release of endospores from mature sporangia. Unfortunately, his pioneering observations were ignored and not duplicated. Our observations confirm Ashworth’s belief. We found that endospore release was initiated when mature sporangia come in 102 contact with water. Following the addition of water, the sporangia were visibly stimulated and their endospore were released through an exit pore located in their cell walls. The finding of a pore in the early stages of sporangial development, suggests that the site of the pore’s formation is probably genetically determined and not a random event. The presence of a pore in mature sporangia has been previously documented (4, 7, 13, 15, 19). However, those reports were based on histological data. Thus, the finding of a pore was always fortuitous and depended on the plane in which the sporangia had been sectioned. Because only few reports of such openings were published, its presence and role were always a subject of debate. Our data, however, indicate that R. seeberi releases its endospores through a pore and that there is only one pore per sporangium. This finding supports Ashworth (6), Grover (9), and Levy et al. (16) who also had observed the release of endospores through a pore in fresh sporangia. Although, Grover (9) and Levy et al. (16) claimed that they had successfully cultured R. seeberi, what they actually observed was the release of endospores from mature sporangia after incubation, and not the product of in vitro multiplication. Their claims have never been substantiated by other investigators. The mechanisms by which water stimulates pore formation and endospore release in R. seeberi are unknown. Ashworth (6) believed that water was absorbed through R. seeben‘s sporangial wall and that this process increased the sporangium’s lntemal pressure. He postulated that the sporangium’s lntemal pressure opened the pore and triggered the release of the endospores. Our study tend to corroborate his observations. Based on our data, we believe that upon contact with water several mechanisms of endospore release are activated. It is possible that at maturity the expression of lytic enzymatic substances may take place at the site of pore formation. This could 103 facilitate the opening of the pore and the discharge of endospores. The stimuli that water exerts on the release of endospores from mature sporangia could explain why most rhinosporidial lesions are located in mucous membranes (nose, mouth) that possess watery secretions. A One intriguing feature of R. seeben‘s sporangia, previous to and after endospore release, was the lntemal rearrangement of its endospores. Our observations showed that in a sporangium stimulated by water well developed endospores were present in the upper half of the sporangium facing the pore, whereas the immature endospores (usually smaller and oval) were observed in the lower half of the sporangium. Although this phenomenon also has been reported by others, their descriptions were based on histopathological and ultrastructural studies (7, 13, 15, 19). Bader and Grueber (7) first called attention to the presence of a "germinative zone" in the lower half of the sporangium. They reported that this "germinative zone" contained small and oval endospores 1.5 to 2.0 um in diameter that appeared to be actively formed in that region. These investigators believed that this was an important region of mature sporangia neglected by previous authors. A similar observation was published later by Savino and Margo (17). They also reported that "newly oval immature endospores seem to arise from an electron lucent inner layer in the sporangial cell wall”. Our in vitro study confirms the formation of new endospores from this complex sector of the sporangia. We believe that the term "germinative zone" is appropriate and should be accepted and used to describe this sporangial region. To confirm that the endospores were actively formed in the “germinative zone” during endospore release, we targeted several immature endospores near this region, and followed their movement toward the apical pore. To our surprise, the immature endospores increased in size as they approached the center of the sporangium and by the time they reached the pore they had matured. 104 Meanwhile, more immature endospores had developed in the ”germinative zone". This observation supports the germinative zone’s ability to produce endospores and confirmed previous observations on this fascinating sporangial region (7, 17). I Recently, a mucoid inner sporangial layer was proven to possess antigenic properties (11). It was found that the antibodies, present in individuals with rhinosporidiosis, detected a mucoid layer beneath the sporangium’s cell wall. Because this layer had immunogenic properties, it was suggested that it may play a role in the immunology of rhinosporidiosis. We could not determine if the “germinative zone” was the same antigenic region detected by Herr at al. (11). However, both substances are located beneath the cell wall. According to these investigators (11), the antigenic material was always detected near mature endospores. This suggests that the antigenic substance is near the pore. Moreover, the mucous layer near the pore had different electrolucent properties than the one in the germinative zone. Based on Herr et al’s (11) observations and the data in this study, we speculate that perhaps the upper electrolucent layer is important in the immunology of rhinosporidiosis, whereas the “germinative zone” is involved in the production of endospores during the process of endospore release. All in all our studies support the concept that R. seeberi is a hydrophilic microorganism (3). This concept is in agreement with the molecular data that clustered R. seeberi together with other aquatic pathogens (12), and strongly supports the hypothesis that aqueous habitats are the prime source of rhinosporidial infections. 105 REFERENCES 10. 11. Ahluwalia, K. B., N. Maheshwari, and R. C. Deka. 1997. Rhinosporidiosis: A study that resolves etiologic controversies. Amer. J. Rhinol. 1 1 :479-483. Ahluwalia, K. B. 1992. New interpretations in rhinosporidiosis, enigmatic disease of the last nine decades. J. Submicrosc. Cytol. Pathol. 24:109- 1 14. Ajello, L. 1998. Ecology and epidemiology of hydrophilic infectious fungi and parafungi of medical mycologycal importance: A new category of pathogens, pp. 67-73, Vol. 4, chap. 4. In L. Ajello, R..J Hay, (Eds), Topley & Wilson?s Microbiology and Microbial Infections, 9th ed. Arnold, London. Apple, D.J. 1983. “Papillome”der conjunktiva bendingt durch rhinosporidiose. Fortschr. Ophthalmol. 79:571-574. Arseculeratne, S. N., and L. Ajello. 1998. Rhinosporidium seeberi. pp. 596-615, Vol. 4 Chap. 31. In: L. Ajello, R.J. Hay, (Eds.,) Topley& Wilson?s Microbiology and Microbial Infections, 9th ed. Arnold, London. Ashworth, J. H. 1923. On Rhinosporidium seeberi (Wemicke, 1903) with special reference to its sporulation and affinities. Trans. Roy. Soc. Edinb. 53:301-342. Bader, G. and H.L.E. Grueber. 1970. Histochemical studies of Rhinosporidium seeberi. Virch. Arch. Abt A Path. Anat. 350276-86. Chuan-Teh, E. and M. Kannan-Kutty. 1975. Rhinosporidium seeberi: spherules and their significance Pathol. 7:133-137. Grover, S. 1970. Rhinosporidium seeberi; A preliminary study of the morphology and life cycle. Sabouraudia 7:249-251. Hanaichi,T., T. Sato, T. lwamoto, J. Malavasi-Yamashiro, M. Hoshlno, and N. Herr, R.A., L. Mendoza, S.N. Arseculeratne, and L. Ajello. 1998. lmmunolocalization of an endogenous antigenic material of Rhinosporidium seeberi expressed only during mature sporangial development. FEMS lmmunol. Med. Microbiol. 23:205-212. 106 12. 13. 14. 15. 16. 17. 18. 19. Herr, R.A., L. Ajello, J.W. Taylor, S.N Arseculeratne, and L. Mendoza. 1999. Phylogenetic analysis of Rhinosporidium seeben’s 188 small subunit ribosomal DNA groups this pathogen among members of the protoctistan Mesomycetozoa clade. J. Clin. Microbiol. 37: Kannan-Kutty, M. and E. Chuan-Teh. 19.74. Rhinosporidium seeberi: An electron microscopic study of its life cycle. Pathol. 6:63-70. Kennedy, F.A., R.R. Buggage, and L. Ajello. 1995. Rhinosporidiosis: a description of an unprecedented outbreak in captive swans (Cygnus spp) and a proposal for revision of the ontogenic nomenclature of Rhinosporidium seeberi. J. Med. Vet. Mycol. 33: 1 57-165. 1 994. Kwon-Chung, K.J. and J.E. Bennett. 1992. Coccidioidomycosis. In: Medical Mycology. Lea and Febiger, Philadelphia, pp 356-396. Levy, M.G., D.J., Meuten, and EB. Breitschwerdt. 1986. Cultivation of Rhinosporidium seeberi in vitro: Interaction with epithelial cells. Science 234:474-476. Sabino, D.F. and CE. Margo. 1983. Conjuntival rhinosporidiosis. Light and electron microscopic study. Ophthalmol. 90:1482-1489. Seeber, G. R. 1900. Un nuevo esporozuario parasito del hombre. Dos casos encontrados en polypos nasales. Thesis, Universidad Nacional, Argentina. Imprenta Libreria Boullosa, Buenos Aires, Argentina. Thianprasit, M. and K. Thagemgpol. 1989. Rhinosporidiosis. In: Current Topics in Med. Mycol. 3:64-85., New York, Springer-Verlag. 107 108 Bibliography 109 Bibliography Agrawal, S., Sharma, K. D., Shrivastava, J. B. 1959. Generalized rhinosporidiosis with visceral involvement: report of a case. Arch. Derrnatol. 80: 22-26. Ahluwalia, K. B., N. Maheshwari, and R. C. Deka. 1997. Rhinosporidiosis: A study that resolves etiologic controversies. Amer. J. Rhinol. 1 1 :479-483. Ahluwalia, K. B. 1992. New interpretations in rhinosporidiosis, enigmatic disease of the last nine decades. J. Submicrosc. Cytol. Pathol. 242109-114. Angunawela, P., DeTissera, A., and Dissanalke, A. S. 1999. Rhinosporidiosis presenting with two soft tissue tumors followed by dissemination. Pathology. 31:57-58. Ajello, L. 1998. Ecology and epidemiology of hydrophilic infectious fungi and parafungi of medical mycologycal importance: A new category of pathogens, p. 67-73, Vol. 4, chap. 4. In L. Ajello, R.J. Hay, (Eds), Topley & Wilson’s Microbiology and Microbial Infections, 9th ed. Arnold, London. Arsecularatne, S. N., and L. Ajello. 1998. Rhinosporidium seeberi. p. 596- 615, Vol. 4 Chap. 31. In L. Ajello, R.J. Hay, (Eds.), Topley & Wilson’s Microbiology and Microbial Infections, 9th ed. Arnold, London. Ashworth, J. H. 1923. On Rhinosporidium seeberi (Wemicke, 1903) with special reference to its sporulation and affinities. Trans. Roy. Soc. Edin. 53:301-342. Broz, O. and M. Privora. 1951. Two skin parasites of Rana temporan'a: Dermocystidium ranae Guyenot and Naville, and Dermospon'dium granulosum n.sp. Parasitology 42:65-69. Carini, A. 1940. Sobre um parasito semlhante ao Rhinosporidium, encontrado em quistos da pele de uma hyla. Arquivos lnstituto Biologico, Sao Paulo, Brazil, 11 :93-96. Cervinka, 8., J. Vitovec, J. Lom, J. Hoska, and F. Kubu. 1974. Dermocystidiosis: A gill disease of the carp due to Dermocystidium cypn'ni n.sp. J. Fish Biol. 6:689-699. Chatterjee, P. K., Khatua, C. R., Chatterjee, S. N., et al. 1977. Recurrent multiple rhinosporidiosis with osteolytic lesions in hand and foot: a case report. J. Laryngol. Otol. 91: 727-734. 110 Davidson, W. R., Nettles, V. F. 1977. Rhinosporidiosis in a wood duck. JAVMA. 171: 989-990. Dodge, C. W. 1935. Rhinosporidium. In Medical Mycology. Fungous Diseases of Men and other Mammals. p. 151-152. CV. Mosby Co., St. Louis. Dunkerly, J. S. 1914. Dermocystidium pusula Perez, parasitic on trutta fario. Zool. Anz. 44:179-182. Dykova, I and J. Lom. 1992. New evidence of fungal nature of Dermocystidium koi 1 950. J. Appl. Ichthyol. 8: 1 80-185. Elston, R. A., L. Harrell, and M. T. Wilkinson. 1986. Isolation and in vitro characteristics of Chinook salmon (Oncorhynchus tshawytscha) rosette agent. Aquaculture 56:1-21. Gargas, A. and P. T. DcPricst. 1996. A nomenclature for fungal PCR primers with examples from intron-containing SSU rDNA. Mycologia 88:745-748. Grover, S. 1970. Rhinosporidium seeberi: A preliminary study of the morphology and life cycle. Sabouraudia. 7: 249-51. Hanaichi,T., T. Sato, T. lwamoto, J. Malavasi-Yamashiro, M. Hoshlno, and N. Mlzuno, N. 1986. A stable lead by modification of Sato’s method. J. Electron. Microsc. 35:304-306. Harrell, L. W., R. A. Elston, T. M. Scott, and M. J. Wilkinson. 1986. A significant new systemic disease of Net-Pen reared Chinook salmon (Oncorhynchus tshawytscha) brood stock. Aquaculture 55:249-262. Hedrick, R. P., C. S. Friedman, and J. Modin. 1989. Systemic infections in atlantic salmon Salmo salar with a Dermocystidium-like species. Dis. Aquat. Org. 7:171-177 Herr, R. A., Mendoza, L., Arseculeratne, S. N., Ajello, L. 1999. lmmunolocalization of an endogenous antigenic material of Rhinosporidium seeberi expressed only during mature sporangial development. FEMS lmm. And Med. Microbiol. 23: 205-212. Ho, M. 8., Tay, B. K., 1986. Disseminated rhinosporidiosis. Ann. Acad. Med. Singapore. 15: 80-83. lssakainen, J., J. Jalava, E. Eerola, and C. K. Campbell. 1997. Related ness of Pseudallescheria, Scedosporium, and Graphium pro parte 111 based on SSU rDNA sequences. J. Med. Vet. Mycol 35:389-398. Jimenez, J. P., Young, D. E., Hough, A. J. 1984. Rhinosporidiosis. A report of two cases from Arkansas. Am. J. Clin. Pathol. 82: 611-615. Kannan-Kutty, M., Teh, E. C. 1974. Rhinspon'dium seeberi: An electron microscopic study of its life cycle. Pathology. 6: 63-70. Karunaratne, WAE,. 1964. Rhinosporidiosis in Man. The Athlone Press, London. Khan Ali Afzal, Khaleque, K. A., Huda, M. N. 1969. Rhinosporidiosis of the nose. J. Laryngol. Otol. 83: 461-473. Krishnamoorthy, 8., Sreedharan V.P., et al., 1989. Culture of Rhinosporidium seeberi: Preliminary report. J. Laryngol Otol. 103: 178-80. Kerk, D., A. Gee, M. Standish, P. O. Wainwright, A. 8. Drum, R. A. Elston, and M. L. Sogin. 1995. The rosette agent of Chinook salmon (Oncorhynchus tshawytscha) is close related to choanoflagellates, as determined by the phylogenetic analysis of its small ribosomal subunit RNA. Marine Biol. 122:187-192. Kwon-Chung, K. J. 1994. Phylogenetic spectrum of fungi that are pathogenic to humans. Clinical Infectious Diseases. 19(Suppl 1): 81-7. Laveran, A. and A. Pettlt. 1910. Sur une epizootie des truites. CR. Acad. Sci. Paris 151 :421-423. Levy, M.G., D. J. Meutem, and E. B. Breitschwerdt. 1986. Cultivation of Rhinosporidium seeberi in vitro: Interaction with epithelial cells. Science 234:474-476 Mahakrisnan, A., Rajasekaram, V., Pandlan, P. J. 1981. Disseminated cutaneous rhinosporidiosis treated with dapsone. Trop. Geogr. Med. 33: 189- 192. Maniatis, T., Fristch, E. F., Sambrook, J. 1982. Molecular cloning: A laboratory manual. Cold Spring Harbor laboratory. Margulis, L. and Schwartz, K. V. 1998. Five Kingdoms an illustrated guide to the phyla of life on earth. 3rd ed. W. H. Freeman and Company. Mosier, D. A., Creed, J. E. 1984. Rhinosporidiosis in dog. JAVMA. 185: 1009-1010. 112 Olsen, G. J. 1987. The earliest phylogenetic branchings: compairing rRNA- based evolutionary trees inferred with various techniques. Cold Spring Harbor Symp. Quant. Biol. 52: 825-838. Olsen, G. J., and Woese, C. R. 1989. A brief note concerning archaebacterial phylogeny. Canadian Journal of Microbiology. 35: 119-123. Ragan, M. A., C. L. Goggin, R. J. Cawthom, L. Cerenius, A. V. C. Jamieson, S. M. Plourdes, T. G. Tand, K. Soderhall, and R. R. Gutell. 1996. A novel clade of protistan parasites near the animal-fungal divergence. Proc. Natl. Acad. Sci. 93:11907-11912. Rajam, R. V., Viswanathan, G. 8., Rao, A. R., et al. 1955. Rhinosporidiosis: a study with report of a fatal case of systemic dissemination. Indian J. Surg. 17: 269-298. Raddy, D., Lakshminarayana, C. S. 1962. Investigation into transmission, growth, and serology in rhinosporidiosis. Indian J. Med. Res. 50: 363-370. Rippon, J. W. 1982. Medical Mycology; the pathogenic fungi and the pathogenic actinomycetes, in Rhinosporidiosis. Saunders, Philadelphia, 325- 334. Savino, D. F., Margo, C. E. 1983. Conjunctival Rhinosporidiosis light and electron microscopic study. Opthalmology. 91: 1482-1489. Satyanarayana, C. 1966. Clinical Surgery. ButterWorths, London. 143-152. Seeber, G. R. 1900. Un nuevo esporozuario parasito del hombre. Dos casos encontrados en polypos nasales. Thesis, Imprenta Libreria Boullosa, Buenos Aires, Argentina. Siddall, M. E., D. S. Martin, D. Bridge, 8. S. Desser, and D. K. Cone. 1995. The demise of a phylum of protists: Phylogeny of myxozoa and other parasitic cnidaria. J. Parasitol. 81:961-967. Smith, H. A., Frankson, M. C. 1961. Rhinosporidiosis in a Texas horse. The Southwestern Veterinarian. FALL 1961: 22-24. Smothers, J. F., C. D. von Dholen, L. H. Smith Jr., R. D. Spall. 1994. Molecular evidence that the myxozoan protists are metazoans. Science 265:1719—1721. Spanggaard, B., H. H. Huss, and J. Brescianl. 1995. Morphology of Ichthyophonus hoferi assessed by light and scanning electron microscopy. J. Fish Dis. 18:567-577. 113 Spanggaard, B., P. Skounoe, L. Rossen, and J. W. Taylor. 1996. Pylogenetic relationships of the intracellular fish pathogens Ichthyophonus hoferi, fungi, choanoflagellates, and the rosette agent. Marine. Biol. 26:109- 115. Thomas, T., Gopinath, N., Betts, R. H. 1956. Rhinosporidiosis of the bronchus. Br. J. Surg. 44: 316-319. Vanbreuseghem, R. 1973. Ultrastructure of Rhinosporidium seeberi. Int. J. of Dermatology 12: 20-28. Vukovic, Z., Bobic-Radovanovic, A., Latkovic, Z., Radovanovic, Z. 1995. An epidemiological investigation of the first outbreak of rhinosporidiosis in Europe. Journal of Tropical Medicine and Hygiene. 98:333-337. Wainright, P. O., G. Hinkle, M. L. Sogin, and S. K. Stickel. 1993. Monophyletic origins of the metazoa: An evolutionary link with fungi. Science 260:340-342. Woodard, B. H., Hudson, J. 1984. Rhinosporidiosis: Ultrastructural study of an infection in South Carolina. South. Med. J. 77: 1587-1588. Wright, R. E. 1922. Rhinosporidium kenealyi of the conjunctiva. Indian Med. Gaz. 57: 82-83. 114 IIIIIIIIIIIIIIIIIIIIIII Mill Hill 3 1293 02487 3