Specimens

We obtained a sample of frozen, minced, infected canine nasal polyp in tissue culture media that had been used for the limited propagation of R. seeberi in cell culture (6). After thawing and a 1-minute centrifugation at 500 x g, approximately 0.2 g of the tissue pellet was digested by mechanical disruption and the DNA was purified by adsorption to glass milk in the presence of a chaotropic agent, according to the manufacturer's instructions (Fast Prep, Bio 101, Vista, CA). The DNA was resuspended in 100µL of 10 mM Tris, 1 mM EDTA buffer at pH 8.5.

Three blocks of fixed, paraffin-embedded nasal polyps from unrelated patients with histologically confirmed rhinosporidiosis were obtained. One tissue sample came from a patient born in southern Asia but living in the United States, and two samples came from patients living in Spain. Twenty-three blocks of fixed, paraffin-embedded nasal polyps from patients without rhinosporidiosis were obtained from 12 consecutive patients at the Palo Alto Veterans Affairs (VA) hospital who had undergone nasal polypectomy. Two 25-µm sections were cut from each block, and the sections were deparaffinized and digested (5). DNA from the digests was then purified by the Isoquick method (ORCA Research, Inc., Bothell, WA), and the DNA was resuspended in 25µL of Tris/EDTA buffer. Sections from a block of fixed, paraffin-embedded human lymph node (histologically normal) were also used in some experiments as negative controls. Tissue sections of C. immitis in bone were obtained from a patient with disseminated coccidioidomycosis at the Palo Alto VA hospital and used for fluorescence in situ hybridization (FISH). Samples of Rosette agent and Dermocystidium salmonis DNA were obtained from the Bodega Marine Laboratory of the University of California, Davis.

Fungal specimens in culture were obtained from laboratories at Stanford University and the Palo Alto VA Health Care System. A cotton swab was used to transfer fungal cells from agar into a 1.5-mL microfuge tube containing 0.5 mL digestion buffer (9) and 0.1 mL glass beads. Samples were incubated at 55°C overnight, and the proteinase k was inactivated at 95°C for 10 minutes and then subjected to two freeze-thaw cycles by immersing tubes in a dry ice-isopropranol bath followed by vortex mixing.

Consensus Polymerase Chain Reaction (PCR) of the 18S rRNA Gene

Broad-range fungal PCR primers were designed from a database of >4,000 small subunit rDNA sequences, with the ARB software package (Technical University, Munich, Germany). Primers were selected that would anneal to most fungal and some protist 18S rDNA but not to 18S rDNA from the chordata (F1-fw, F2-rev, F3-rev) (Table). When the specificity of the primer pairs was tested by using human lymphocyte DNA, no amplification was observed (data not shown). The broad range of primers F1-fw/F2-rev was tested by using DNA from several diverse fungi. Amplification products of the expected size were produced by using DNA from Aspergillus oryzae, Alternaria alternata, Candida albicans, Saccharomyces cerevisiae, Trichyphyton rubrum, Panus rudis, Neurospora crassa, Fusarium solani, Beauveria bassiana, Flammulina velutipes, Gibberella zeae, and Pleurotus ostreatus (data not shown).

PCR consisted of 40 cycles of amplification on a Perkin-Elmer GeneAmp 2400 thermal cycler. After an initial activation of Taq gold at 94°C for 10 minutes, each cycle consisted of 30 seconds of melting at 94°C, 30 seconds of annealing at 56°C, and 30 seconds of extension at 72°C. The last cycle was followed by an extension step at 72°C for 7 minutes. Amplification products were detected by electrophoresis on 2% agarose gels stained with ethidium bromide and visualized with a UV transilluminator.

On the basis of the sequences obtained by consensus PCR with primers F1-fw/F2-rev (~500 bp) and F1-fw/F3-rev (~1000 bp), primers Dermo-fw and Dermo-rev were designed (Table) and used in a PCR to amplify a more complete 18S rDNA sequence of R. seeberi.

Rhinosporidium-Specific PCR

A pair of PCR primers was designed from unique regions of the R. seeberi 18S rRNA gene sequence (Rhino-fw and Rhino-rev) (Table). These primers were used in a 50-µL PCR as described, except that AmpliTaq DNA polymerase (PE-ABI) was used at 1 unit per reaction, no dimethyl sulfoxide was added, 50 cycles of PCR were run with a 3-minute pre-melt at 94°C, and the annealing temperature was 55°C. To each 50-µL PCR reaction, 1µL or 5µL of purified DNA were added.

ß-Globin PCR

ß-globin PCR was performed on control tissues as described previously (9).

DNA Cloning, Sequencing, and Phylogenetic Analysis

Amplification products were cloned by using the Topo-TA cloning kit (Invitrogen, Carlsbad, CA), and three clones were sequenced. Each clone consisted of 1,750 bp of 18S rDNA. Priming sequences were removed for further analysis, yielding 1,699 bp of meaningful sequence. DNA sequencing was performed as described (10). A consensus sequence from the three clones was made to correct for any Taq polymerase incorporation errors. The 18S rDNA primers (Table) were used as sequencing primers.

The R. seeberi 18S rDNA sequence was aligned by using the automated aligner of the ARB software package. Ambiguously and incorrectly aligned positions were manually aligned on the basis of the conserved primary sequence and secondary structure. The phylogenetic relationship of R. seeberi to other eukaryotes was inferred from 1,350 unambiguously aligned (masked) positions with a maximum-likelihood algorithm (11,12), on the basis of a previously aligned dataset of the DRIPs clade (named after the organisms Dermocystidium, the Rosette agent, Ichthyophonus, and Psorospermium) (13). The dataset was used to empirically determine nucleotide frequencies and instantaneous substitution rates with the restriction of a 2:1 transition to transversion ratio. The organisms used in our tree and the accession numbers for their small subunit rRNA sequences include Artemia salina (X01723), Xenopus laevis (X04025), Mytilus edulis (L24489), Tripedalia cystophora (L10829), Microciona prolifera (L10825), Diaphanoeca grandis (L10824), Rosette agent (L29455), R. seeberi (AF158369), Dermocystidium species (U21336), Dermocystidium salmonis (U21337), Psorospermium haeckelii (U33180), Ichthyophonus hoferi (D14358), Aspergillus fumagatus (M60300), Chytridium confervae (M59758), Mucor racemosus (X54863), Acanthamoeba castellanii (U07413), Zamia pumila (M20017), Porphyra spiralis (L26177), Lagenidium giganteum (X54266), Labyrinthuloides minuta (L27634), Perkinsus marinus (X75762), Sarcocystis muris (M64244). The tree topology was confirmed by using a neighbor-joining algorithm with Jukes-Cantor corrected distance values and a maximum-parsimony algorithm (ARB). The nucleotide sequence for the partial 18S rRNA gene of R. seeberi has been deposited in GenBank (accession number AF158369).

Fluorescence in Situ Hybridization (FISH)

Tissue sections on slides were dewaxed by immersion in 99% octane (Sigma, St. Louis, MO). Samples subjected to FISH included R. seeberi-infected human nasal polyps, C. immitis-infected bone, a Rosette agent-infected cell line, and smears of C. albicans. The Rhinosporidium probe was based on the Rhino-rev 18S rDNA primer and was biotinylated at both the 5' and 3' ends (Table). The control probe, which consisted of the complement of the Rhinosporidium probe, was also biotinylated at both ends. To each slide, 50 ng of biotinylated probe in 30 µL of hybridization buffer was added. Cover slips were placed, and the slides were incubated at 40°C overnight in a humid chamber. The hybridization buffer consisted of 10% dextran, 0.2% bovine serum albumin, and 0.01% polyadenosine, in 5X SET buffer; the 25X SET buffer consisted of 3.75M sodium chloride, 25 mM EDTA, and 0.5M Tris at pH 7.8. Cover slips were removed by immersion in 5X SET buffer at 4°C, and the slides were washed for 10 minutes per cycle, twice in 0.2X SET buffer at 25°C and once at 40°C. The slides were then subjected to tyramide signal amplification according to the manufacturer's instructions (TSA indirect, NEN Life Sciences, Boston, MA). Cy5-streptavidin (Amersham, Piscataway, NJ) at 1 mg/mL was diluted 1:500 and added to the slides for fluorescence signal detection. Tissue sections were visualized on a Bio Rad confocal microscope at 200X magnification after the application of 15-20µL of Vectashield mountant (Sigma) and a cover slip.

Electron Microscopy

A portion of formalin-fixed, paraffin-embedded nasal polyp from a patient with rhinosporidiosis was removed from the block, dewaxed with xylene, rehydrated with ethanol, post-stained with 1.5% osmium tetroxide, then dehydrated with ethanol, transferred to propylene oxide followed by Epon 12 resin, heat-catalyzed at 65°C, and ultrasectioned at 50 nm. The grid-mounted sections were then serially stained with lead hydroxide and uranyl acetate and examined with a Phillips 201 electron microscope at 75KV.

Phylogenetic Classification of R. seeberi Inferred from the 18S rRNA Gene

Figure 2

Figure 2. Phylogeny of Rhinosporidium seeberi and the DRIPs clade of protists (Ichthyosporea). A. Phylogenetic tree inferred from the 18S rDNA sequences of R. seeberi and other selected eukaryotes by using a maximum...

Consensus PCR of the 18S rRNA gene with DNA from a digest of an R. seeberi-infected canine nasal polyp produced amplification products of the expected size visible on gel electrophoresis (primer pairs F1-fw/F2-rev = ~500 bp and F1-fw/F3-rev = ~1000 bp) (data not shown). No amplification product was detected by using control tissue and reagents. On the basis of the initial phylogenetic assessment of these sequences, primers Dermo-fw and Dermo-rev were designed for amplification of a more complete portion of the R. seeberi 18S rRNA gene. Our phylogenetic analysis of this gene suggests that R. seeberi is a member of the DRIPs clade of aquatic protistan parasites (Figure 2). The nearest evolutionary neighbors of R. seeberi for which a sequence is available are members of the Dermocystidium genus, which infect salmon and trout.

Development and Use of a Rhinosporidium-Specific PCR Assay

A PCR assay specific for R. seeberi was developed. Primers were created by aligning 18S rDNA sequences from R. seeberi, members of the DRIPs clade, Saccharomyces cerevisiae, and humans. The Rhinosporidium primers (Rhino-fw, Rhino-rev) each have three nucleotide mismatches with the sequences from the nearest phylogenetic relatives in the Dermocystidium genus and multiple other mismatches with fungal and human 18S rDNA sequences. An assay sensitivity of 1-10 gene copies was demonstrated by using a dilution series of cloned R. seeberi 18S rDNA. The specificity of the assay was assessed with DNA from human lymphocytes, S. cerevisiae, D. salmonis, and the Rosette agent. No amplification was detected when these DNA samples were used in the specific PCR assay, although product was amplified from these samples with either ß-globin primers (lymphocytes) or broad-range 18S rDNA primers (F1-fw/F2-rev) (Table).

Figure 3

Figure 3. Agarose gel electrophoresis of Rhinosporidium-specific PCR products. The specific amplification product is 377 bp. No amplification product is seen in the negative control samples consisting of water (reagent-only control), digestion buffer...

The original DNA sample from the infected canine nasal polyp yielded a product of the expected size with the Rhinosporidium-specific PCR assay (Figure 3). Purified DNA from the tissue blocks of human nasal polyps resected from three patients with rhinosporidiosis also yielded positive results in this PCR assay, with visible bands seen on gel electrophoresis (RS 1-3) (Figure 3). Direct sequencing of these PCR products demonstrated complete identity over the 377 bp with the cloned sequence from the canine polyp. DNA samples from 23 nasal polyp specimens resected from 12 patients without rhinosporidiosis were subjected to both Rhinosporidium-specific and ß-globin PCR assays. These uninfected polyps failed to yield visible amplicons after gel electrophoresis of the Rhinosporidium-specific PCR reactions (data not shown). However, all these samples yielded visible amplification products after ß-globin PCR, demonstrating that amplifiable DNA was present, without substantial PCR inhibition. These results confirmed that the presence of the putative R. seeberi 18S rDNA sequence correlated with the presence of disease (rhinosporidiosis).

Fluorescence in Situ Hybridization

Figure 4

Figure 4. An oligonucleotide probe complementary to a unique region of the Rhinosporidium seeberi 18S rRNA sequence localizes to visible organisms in human nasal tissue by using fluorescence in situ hybridization. Confocal micrographs...

We sought to determine by using FISH if the R. seeberi 18S rDNA sequence was linked to visible pathology in tissue. The Rhinosporidium 18S rRNA probe but not the control probe bound to R. seeberi organisms in tissue, providing further evidence that the putative R. seeberi 18S rRNA sequence is present in R. seeberi organisms (Figure 4). To test the specificity of the hybridization, smears of the fungus C. albicans, cells infected with the Rosette agent, and tissue sections of C. immitis were subjected to FISH. No specific hybridization to these organisms was detected by using the Rhinosporidium 18S rRNA probe compared with the control probe (data not shown).

Electron Microscopy

Figure 5

Figure 5. Transmission electron micrograph of a mitochondrion from Rhinosporidium seeberi. The cristae of this mitochondrion (arrows) have tubulovesicular morphology. Magnification 195,000X.

Transmission electron micrographs were taken of R. seeberi trophocytes in a human nasal polyp. Multiple mitochondria were visualized, showing that the cristae had tubulovesicular morphology (Figure 5), unlike the flat cristae found in fungi.