Abstract
The in vitro susceptibilities of 66 molecularly identified strains of the Mucorales to eight antifungals (amphotericin B, terbinafine, itraconazole, posaconazole, voriconazole, caspofungin, micafungin, and 5-fluorocytosine) were tested. Molecular phylogeny was reconstructed based on the nuclear ribosomal large subunit to reveal taxon-specific susceptibility profiles. The impressive phylogenetic diversity of the Mucorales was reflected in susceptibilities differing at family, genus, and species levels. Amphotericin B was the most active drug, though somewhat less against Rhizopus and Cunninghamella species. Posaconazole was the second most effective antifungal agent but showed reduced activity in Mucor and Cunninghamella strains, while voriconazole lacked in vitro activity for most strains. Genera attributed to the Mucoraceae exhibited a wide range of MICs for posaconazole, itraconazole, and terbinafine and included resistant strains. Cunninghamella also comprised strains resistant to all azoles tested but was fully susceptible to terbinafine. In contrast, the Lichtheimiaceae completely lacked strains with reduced susceptibility for these antifungals. Syncephalastrum species exhibited susceptibility profiles similar to those of the Lichtheimiaceae. Mucor species were more resistant to azoles than Rhizopus species. Species-specific responses were obtained for terbinafine where only Rhizopus arrhizus and Mucor circinelloides were resistant. Complete or vast resistance was observed for 5-fluorocytosine, caspofungin, and micafungin. Intraspecific variability of in vitro susceptibility was found in all genera tested but was especially high in Mucor and Rhizopus for azoles and terbinafine. Accurate molecular identification of etiologic agents is compulsory to predict therapy outcome. For species of critical genera such as Mucor and Rhizopus, exhibiting high intraspecific variation, susceptibility testing before the onset of therapy is recommended.
INTRODUCTION
The fungal order Mucorales, belonging to a section of lower fungi that until recently was referred to as zygomycetes, constitutes a phylogenetically ancient group of organisms. In the fungal tree of life the group encompasses a number of widely spaced, ancestral lineages. Over time, mutations are hypothesized to have accumulated, which is reflected, e.g., in an immense degree of sequence diversity of evolutionary markers such as the ribosomal operon. By assessment of identical genes, mucoralean species are separated from each other at branches much longer than those of species of more recent fungi, such as Aspergillus or the dermatophytes. As a result, ancestry is difficult to reconstruct, leading to phylogenetic trees with poorly resolved backbones. For similar reasons, the phylum Zygomycota has been abandoned: phylogenetic distances are so large that no taxonomic hierarchy can be constructed and no umbrella group defined that would unite all fungi attributed to the Zygomycota in the classical sense (23).
The opportunistic members of the Mucorales are classified in the families Cunninghamellaceae, Lichtheimiaceae, Mucoraceae, Saksenaceae, and Syncephalastraceae, with the great majority of human infections being caused by members of Mucoraceae and Lichtheimiaceae. In molecular phylogenetic analyses (31, 43), the genus Rhizomucor was positioned outside the Mucoraceae, and in the present article the Index Fungorum (http://www.indexfungorum.org) is followed, classifying the genus in the Lichtheimiaceae.
Infections generally occur in severely debilitated patients and are acute, destructive, and with a rapid course and fatal outcome (14, 37). In general, different types of underlying conditions predispose for different types of infection. Major skin abrasion and burn wounds may lead to erosive subcutaneous infection. Rhinocerebral and pulmonary infection are linked to ketoacidotic diabetes and severe neutropenia, respectively, while immunosuppression and prolonged deferoxamine therapy predispose for disseminated infection (11). Chronic disorders observed in individuals without severe immune or metabolic dysfunction are exceptional cases (25). Also, renal mucormycosis tends to occur in immunocompetent individuals (21, 27).
Given the enormous phylogenetic diversity of the Mucorales (31, 43), it is remarkable that frequent case reports appear referring to the etiologic agent without proper species identification (e.g., references 19 and 26). Practical reasons for this are the difficulties of cultivating these fungi from biopsy samples (22, 34) and of differentiating zygomycete species by classical mycological techniques in clinical microbiology laboratories. Previous comprehensive studies on susceptibility profiles against antifungal drugs in the Mucorales revealed considerable variation among and within genera and species, as defined either by applying classical parameters (5, 16, 35, 39, 40) or more recently by using molecular taxonomic methods (1, 12). A recent review of in vitro activity of antifungals against zygomycetes was provided by Alastruey-Izquierdo et al. (2).
Application of molecular methods for species identification in Mucorales frequently leads to the unexpected detection of novel sibling species (3, 6, 9). For this reason, and because of the difficulty of morphological species identification, studies on the taxon specificity of susceptibility profiles in this group of fungi need to be preceded by molecular identification of the strains under study (1). Otherwise, in clinical practice, when the only intention is to determine the most appropriate antifungal as quickly as possible, susceptibility tests could also be performed directly. We chose the internal transcribed spacer (ITS) region for taxonomy at the species level because it earlier has been shown to be the species marker of choice in Mucorales (36).
Classification of Mucorales above the species level is in a state of flux, since molecular phylogenetic analyses found polyphyly of the majority of morphology-based families and genera (31, 41). The D1/D2 region of the nuclear ribosomal large subunit (LSU) was chosen to reconstruct phylogeny because it could be sequenced directly, while all protein-encoding genes tested revealed paralogs in numerous species (3). Furthermore, the LSU is alignable over the entire order. A robust molecular phylogenetic hypothesis is necessary to address the main question of this study: do phylogenetic taxa (species, genera, and families) of the Mucorales possess more or less characteristic susceptibility profiles?
MATERIALS AND METHODS
Strains.
All isolates used in this study were taken from the reference collection of the CBS-KNAW Fungal Biodiversity Centre (CBS, Utrecht, The Netherlands), from the Institut Pasteur (CNRMA/IP, Paris, France), or from the American Type Culture Collection (ATCC, Manassas, VA). Strains selected for phylogenetic reconstruction represented all relevant taxa of the Mucorales including the clinically relevant species.
Extraction of genomic DNA, amplification, cloning, and sequencing.
Genomic DNA was extracted from 2-day-old malt extract agar (MEA) cultures according to the procedure reported by Möller et al. (28) with several modifications described in detail by Alastruey-Izquierdo et al. (3). DNA segments comprising the complete ITS region and the D1/D2 region of the LSU were amplified using the primer pair V9G (18) and LR3 (41). The PCR mixture (25 μl) contained 0.4 μM each primer, 0.185 mM each deoxynucleoside triphosphate (GC Biotech, Alphen a/d Rijn, The Netherlands), 10× NH4 BioTaq reaction buffer (GC Biotech), a final concentration of 1.5 mM MgCl2, 0.8 U BioTaq DNA polymerase (GC Biotech), and about 20 ng DNA. The cycling conditions included one initial cycle at 94°C for 5 min, followed by 35 cycles of 1 min at 94°C, 1 min at 53°C, and 2 min at 72°C, with one final cycle of 7 min at 72°C. PCRs were performed on a thermal cycler 2720 (Applied Biosystems, Nieuwerkerk a/d IJssel, The Netherlands). Reaction products were analyzed in 1% agarose gels. Both strands of the PCR products were directly cycle sequenced using the BigDye sequencing kit (Applied Biosystems), and the primer set ITS1 and ITS4 (44) for the complete ITS region and NL1 and LR3 for the D1/D2 region of the LSU (30) were used. Cycle-sequencing products were analyzed on an ABI 3730XL automatic sequencer (Applied Biosystems). In a few cases, direct sequencing failed and the PCR products were cloned in the competent Escherichia coli cell line JM109 by using the pGEM-T Easy vector (Promega, Leiden, The Netherlands) as instructed by the manufacturer. Colony PCRs were performed using the primer pair M13f (5′-GTAAAACGACGGCCAGT-3′) and M13r (5′-GGAAACAGCTATGACCATG-3′). Products of colony PCRs were sequenced as described above.
Species identification and phylogenetic analysis.
Consensus sequences were constructed by means of the SeqMan program v.7.2.2. (DNASTAR, Lasergene). The D1/D2 region of the LSU was used to reconstruct the phylogeny of the Mucorales, while the ITS region served as a marker for molecular species recognition. For both markers, sequences were aligned using the server version of the MAFFT program (http://www.ebi.ac.uk/Tools/mafft) and manually corrected in the program Se-Al v2.0a11 (http://tree.bio.ed.ac.uk/software/seal/) (33). Sequences of the ITS spacers were not alignable over the entire order. Therefore, ITS alignments for each studied genus were prepared including sequences of all relevant ex-type strains to ensure an accurate identification. They were rooted by the nearest neighbors according to the work of O'Donnell et al. (31). Phylogenetic relationships were estimated using the maximum likelihood method with the server version of RAxML-VI-HPC v.7.0.0 (38), as implemented on the Cipres portal. The robustness of the trees was estimated by a bootstrap analysis with 1,000 replicates.
In vitro susceptibility testing.
A set of 66 strains was tested (Table 1), including species of Cunninghamella (n = 8), Lichtheimia (syn. Absidia pro parte, Mycocladus) (n = 13), Mucor (n = 12), Rhizomucor (n = 8), Rhizopus (n = 19), and Syncephalastrum (n = 6). In vitro susceptibilities were determined by a broth microdilution technique following the guidelines of the Clinical and Laboratory Standards Institute (CLSI) M38A document (29) with few modifications. RPMI 1640 medium with l-glutamine but without sodium bicarbonate (Sigma-Aldrich, Saint Quentin Fallavier, France) buffered to pH 7.0 with 0.165 M MOPS (morpholinepropanesulfonic acid) (Sigma-Aldrich) was used as the test medium. Isolates were grown on Sabouraud dextrose agar for 7 days at 30°C, and stock spore suspensions were prepared by washing the surface of the slants with sterile saline containing 0.05% Tween 80. Spore suspensions were counted with a hemacytometer and then diluted into RPMI medium to a concentration of 2 × 104 spores/ml (2× final concentration). Pure powders of known potency of amphotericin B (AMB; Sigma-Aldrich, Saint Quentin Fallavier, France), voriconazole (VCZ; Pfizer Central Research, Sandwich, United Kingdom), itraconazole (ITZ; Janssen-Cilag, Issy-les-Moulineaux, France), posaconazole (PCZ; Schering-Plough, Kenilworth, NJ), 5-fluorocytosine (5-FC; Sigma-Aldrich), terbinafine (TBF; Novartis Pharma, Basel, Switzerland), caspofungin (CAS; Merck, Rahway, NJ), and micafungin (MCF; Astellas Pharma, Inc., Ibaraki, Japan) were used.
Table 1.
Strains analyzed with present molecular identification and in vitro susceptibility dataa
| Name | Strain | Source | Country | MIC/MEC (μg/ml) for: |
|||||||
|---|---|---|---|---|---|---|---|---|---|---|---|
| 5-FC | MCF | CAS | TBF | VCZ | ITZ | PCZ | AMB | ||||
| Mucoraceae | |||||||||||
| Mucor | |||||||||||
| M. circinelloides (syn. Rhizomucor regularior) | CBS 384.95 T | Human, face | China | >64 | >4 | >4 | 16 | >8 | 4 | 1 | 0.25 |
| M. circinelloides | CBS 416.77 | Fermenting rice | Unknown | >64 | >4 | >4 | 4 | >8 | 2 | 1 | 0.5 |
| M. circinelloides | CNRMA 03.154 | Human, skin | France | >64 | >4 | >4 | 32 | >8 | 1 | 0.5 | 0.25 |
| M. circinelloides | CNRMA 03.371 | Human, muscle | France | >64 | >4 | >4 | 2 | >8 | 2 | 1 | 0.5 |
| M. circinelloides f. circinelloides | CBS 195.68 NT | Air | Netherlands | >64 | >4 | >4 | 32 | >8 | 1 | 1 | 0.25 |
| M. circinelloides | IP 1873.89 | Human, feces | France | >64 | >4 | >4 | 32 | >8 | 2 | 1 | 0.25 |
| M. circinelloides | CNRMA 04.805 | Human, muscle | France | >64 | >4 | >4 | 2 | >8 | 2 | 2 | 0.5 |
| M. ramosissimus | CBS 135.65 NT | Human, nasal lesion | Uruguay | >64 | >4 | >4 | 2 | >8 | 1 | 1 | 0.25 |
| M. racemosus f. racemosus | CBS 260.68 T | Unknown | Switzerland | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.125 | 0.5 |
| M. hiemalis f. hiemalis | CBS 201.65 NT | Unknown | USA | >64 | >4 | >4 | 0.25 | 1 | 0.5 | 0.5 | 0.5 |
| M. irregularis (syn. Rhizomucor variabilis) | CBS 103.93 T | Human, hand | China | >64 | >4 | >4 | 0.5 | >8 | 1 | 1 | 0.5 |
| M. indicus | CNRMA 03.894 | Human, muscle | France | >64 | >4 | >4 | 0.5 | >8 | 1 | 1 | 0.5 |
| Rhizopus | |||||||||||
| R. arrhizus | IP 4.77 | Human, brain | France | >64 | >4 | >4 | 32 | 8 | 0.5 | 0.5 | 1 |
| R. arrhizus | CNRMA 03.413 | Unknown | France | >64 | >4 | >4 | 32 | 4 | 0.5 | 0.25 | 0.5 |
| R. arrhizus | CNRMA 03.410 | Human, sputum | France | >64 | >4 | >4 | 32 | 8 | 1 | 0.5 | 1 |
| R. arrhizus | CNRMA 03.412 | Human, sputum | France | >64 | >4 | >4 | 1 | 4 | 0.5 | 0.125 | 1 |
| R. arrhizus | CBS 112.07 T | Unknown | Netherlands | >64 | >4 | >4 | 32 | 8 | 0.5 | 0.5 | 1 |
| R. arrhizus | CNRMA 03.411 | Human, sputum | France | >64 | >4 | >4 | 4 | 8 | 0.5 | 0.25 | 1 |
| R. arrhizus | CNRMA 04.48 | Human, skin | France | >64 | >4 | >4 | 32 | 8 | 0.5 | 0.25 | 2 |
| R. arrhizus | CNRMA 04.160 | Human, sputum | France | >64 | >4 | >4 | 16 | 8 | 0.5 | 0.25 | 1 |
| R. arrhizus | CNRMA 03.253 | Human, lung | France | >64 | >4 | >4 | 4 | 4 | 0.25 | 0.03 | 1 |
| R. arrhizus | CNRMA 03.395 | Human, skin | France | >64 | >4 | >4 | 32 | 4 | 0.5 | 0.25 | 1 |
| R. arrhizus | CNRMA 03.375 | Human, sinus | France | >64 | >4 | >4 | 32 | >8 | 1 | 0.5 | 2 |
| R. arrhizus | CNRMA 03.909 | Human, sinus | France | >64 | >4 | >4 | 8 | 4 | 0.5 | 0.25 | 0.5 |
| R. arrhizus | IP 1443.83 | Unknown | Unknown | >64 | >4 | >4 | 2 | 4 | 0.125 | 0.5 | 0.5 |
| R. arrhizus | CNRMA 03.918 | Human, lung | France | >64 | >4 | >4 | 32 | 8 | 1 | 0.5 | 1 |
| R. microsporus var. azygosporus | CBS 357.93 T | Tempeh | Indonesia | >64 | >4 | >4 | 0.125 | 4 | 0.5 | 0.5 | 1 |
| R. microsporus var. rhizopodiformis | CNRMA 04.1469 | Human | France | >64 | >4 | >4 | 0.125 | 8 | 4 | 2 | 2 |
| R. microsporus var. chinensis | CBS 631.82 T | Bread | China | >64 | >4 | >4 | 0.25 | 4 | 1 | 0.5 | 1 |
| R. microsporus var. rhizopodiformis | IP 1123.75 | Unknown | Unknown | >64 | >4 | >4 | 0.06 | 4 | 0.5 | 0.5 | 1 |
| R. microsporus var. rhizopodiformis | IP 676.72 | Human, skin | France | >64 | >4 | >4 | 0.125 | 4 | 0.5 | 0.25 | 1 |
| Syncephalastraceae | |||||||||||
| Syncephalastrum | |||||||||||
| S. “racemosum” (species II) | CBS 199.81 | Soil | Kuwait | 64 | >4 | >4 | 0.25 | >8 | 0.5 | 0.25 | 0.06 |
| S. “racemosum” (species II) | CBS 421.63 | Soil | Zaire | >64 | >4 | >4 | 0.25 | >8 | 0.5 | 0.5 | 0.03 |
| S. “racemosum” (species I) | CBS 302.65 | Soil | Brazil | >64 | >4 | >4 | 0.25 | >8 | 0.5 | 1 | 0.25 |
| S. “racemosum” (species I) | CBS 441.59 | Dung of coyote | USA | 64 | >4 | >4 | 0.25 | 8 | 0.5 | 0.25 | 0.06 |
| S. “racemosum” (species I) | CNRMA 03.414 | Human, skin | France | >64 | >4 | >4 | 0.06 | 1 | 0.03 | 0.06 | 0.25 |
| S. “racemosum” (species I) | CBS 370.49 | Air | Indonesia | >64 | >4 | >4 | 0.125 | 8 | 0.25 | 0.5 | 0.06 |
| Lichtheimiaceae | |||||||||||
| Rhizomucor | |||||||||||
| R. pusillus | CBS 354.68 NT | Corn meal | Netherlands | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.25 | 0.5 |
| R. pusillus | CNRMA 04.210 | Human, bone | France | >64 | >4 | >4 | 0.125 | >8 | 0.25 | 0.25 | 0.5 |
| R. pusillus | CNRMA 04.503 | Human, sputum | France | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.5 | 0.5 |
| R. pusillus | IP 1956.90 | Human, bronchia | France | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.25 | 0.5 |
| R. pusillus | ATCC 36606 | Cat, brain | France | >64 | >4 | >4 | 0.06 | 4 | 0.25 | 0.125 | 0.5 |
| R. pusillus | CNRMA 03.1205 | Human, lung | France | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.25 | 0.5 |
| R. pusillus | IP 1127.75 | Unknown | Unknown | >64 | >4 | >4 | 0.06 | 4 | 0.25 | 0.125 | 0.5 |
| R. miehei | CBS 182.67 T | Retting plant | USA | >64 | >4 | >4 | 0.25 | 4 | 0.06 | 0.125 | 0.5 |
| Lichtheimia | CNRMA 03.611 | Human, bronchia | France | >64 | >4 | >4 | 0.06 | 8 | 0.125 | 0.25 | 0.5 |
| L. corymbifera | IP 1129.75 | Air | Morocco | >64 | >4 | >4 | 0.125 | 8 | 0.25 | 0.25 | 0.25 |
| L. corymbifera | CNRMA 03.697 | Human, bone | France | >64 | >4 | >4 | 0.125 | 8 | 0.25 | 0.25 | 0.25 |
| L. corymbifera | IP 1279.81 | Unknown | Unknown | >64 | >4 | >4 | 0.125 | 8 | 0.25 | 0.06 | 0.25 |
| L. corymbifera | CNRMA 04.732 | Human, lung | France | >64 | >4 | >4 | 0.25 | >8 | 0.25 | 0.25 | 0.5 |
| L. corymbifera | IP 1280.81 | Unknown | Unknown | >64 | >4 | >4 | 0.125 | >8 | 0.25 | 0.25 | 0.25 |
| L. ramosa | CBS 100.55 | Unknown | Unknown | >64 | >4 | >4 | 0.12 | 8 | 0.5 | 0.06 | 0.125 |
| L. ramosa | CBS 100.49 | Dung of cow | Indonesia | >64 | >4 | >4 | 0.25 | 2 | 0.03 | 0.06 | 0.06 |
| L. ramosa | CBS 124198 | Culture contaminant | Netherlands | >64 | >4 | >4 | 0.25 | 2 | 0.12 | 0.06 | 0.25 |
| L. ramosa | CBS 582.65 NT | Cacao seeds | Ghana | >64 | >4 | >4 | 0.5 | >8 | 0.5 | 0.25 | 0.125 |
| L. ramosa | CBS 223.78 | Soil | Unknown | >64 | >4 | >4 | 0.25 | >8 | 0.25 | 0.25 | 0.125 |
| L. ornata | CBS 958.68 | Unknown | Unknown | >64 | >4 | >4 | 0.25 | 2 | 0.06 | 0.06 | 0.25 |
| L. ornata | CBS 291.66 T | Dung of bird | India | >64 | >4 | >4 | 0.25 | >8 | 0.12 | 0.06 | 0.25 |
| Cunninghamellaceae | |||||||||||
| Cunninghamella | |||||||||||
| C. bertholletiae | CBS 186.84 | Human, lung | USA | >64 | >4 | >4 | 0.06 | >8 | 16 | 1 | 2 |
| C. bertholletiae | CBS 693.68 | Soil | Yugoslavia | >64 | >4 | >4 | 0.25 | >8 | >64 | 1 | 1 |
| C. bertholletiae | CBS 372.95 | Soil | China | >64 | >4 | >4 | 0.06 | >8 | 16 | 0.5 | 1 |
| C. bertholletiae | CBS 190.84 | Human, heart | USA | >64 | >4 | >4 | 0.06 | >8 | 16 | 0.5 | 2 |
| C. bertholletiae | CBS 191.84 | Human, tibia | USA | 32 | 1 | 4 | 0.06 | >8 | >64 | 0.5 | 2 |
| C. echinulata var. antarctica | CBS 545.75 | Soil | Chile | >64 | >4 | >4 | 0.125 | >8 | 1 | 0.5 | 2 |
| C. echinulata | CBS 766.68 | Unknown | Unknown | >64 | >4 | >4 | 0.06 | >8 | >64 | 1 | 1 |
| C. echinulata | CBS 156.28 | Unknown | Unknown | >64 | 0.5 | 4 | 0.25 | >8 | 16 | 1 | 2 |
Susceptibility data are in μg/ml for 5-fluorocytosine (5-FC), micafungin (MCF), caspofungin (CAS), terbinafine (TBF), voriconazole (VCZ), itraconazole (ITZ), posaconazole (PCZ), and amphotericin B (AMB). Genera are placed in line with the phylogenetic tree of Fig. 1 except for Rhizomucor. Within the genera, strains are arranged according to their position in the trees of Fig. 2. Species boundaries are marked by vertical space. NT, neotype; T, ex-type.
Briefly, microplates containing the antifungal drugs were prepared by batch and stored frozen at −20°C for less than 1 month. MIC endpoints were determined by an automated microplate reader spectrophotometer (Multiscan RC-351; Labsystems Oy, Helsinki, Finland) after 24 h of incubation (an optical density [OD] of >0.15 was required for the drug-free control wells) at 35°C (28°C for Mucor and Rhizomucor). MIC endpoints were defined as ≥50% reduction in growth compared to the drug-free wells, except for AMB, for which a 90% reduction endpoint was used. For echinocandins, minimum effective concentrations (MECs) were determined by reading the microplates with the aid of an inverted microscope. Two reference strains, Candida krusei ATCC 6258 and Candida parapsilosis ATCC 22019, were included in each set of determinations to ensure quality control.
Statistics.
For calculation purposes, high off-scale MICs/MECs were raised to the next higher concentration, while low off-scale MICs were left unchanged. Associations between the phylogenetic order and MICs of the various antifungal agents were tested with the two-tailed Mann-Whitney test using GraphPad InStat version 3.00 (GraphPad Instat Software, Inc., San Diego, CA).
Nucleotide sequence accession numbers.
Sixty LSU sequences (GenBank accession numbers HM849659 to HM849674, HM849676 to HM849707, HM849709, HM849710, and HM849715 to HM849724) and 38 ITS sequences (GenBank accession numbers HM999950 to HM999986 and HQ186304) were newly generated for this study.
RESULTS
The LSU tree (Fig. 1) comprises members of the clinically significant genera Apophysomyces, Cokeromyces, Cunninghamella, Lichtheimia, Mucor, Rhizomucor, Rhizopus, Saksenaea, and Syncephalastrum, all belonging to the order Mucorales. Sequences could be aligned with relative confidence but showed considerable diversity. In contrast, morphological identification of species appeared to be nonpredictive, with extreme differences occurring between neighboring molecular taxa. Ribosomal DNA (rDNA) distances between species belonging to a single genus were considerable. ITS diversity was up to 20% in Lichtheimia and 35% in Mucor, while Rhizopus arrhizus and Rhizopus microsporus deviated 29.8% (data not shown). ITS sequences showed very limited similarity over the entire data set and could not be aligned. Separate trees were therefore constructed for molecular identification (Fig. 2).
Fig 1.
RAxML phylogram based on the D1/D2 region of the LSU. Branches with bootstrap values of 85 or higher are printed in bold. Important branch support values lower than 85 are indicated by numbers near the branches. Ex-type strains are indicated by T (type strain), NT (neotype strain), IT (isotype strain), and HT (holotype strain) following the strain numbers. Asterisks indicate strains included in the susceptibility study. Bars (a to f) mark the genera represented by an ITS dendrogram in Fig. 2.
Fig 2.
RAxML phylograms based on the complete ITS region. Branches with bootstrap values of 85 or higher are printed in bold. In cases of less space or important values lower than 85, branch support values are indicated by numbers near the branches. Ex-type strains are printed in bold and indicated by T (type strain), NT (neotype strain), IT (isotype strain), and HT (holotype strain) following the strain numbers. Asterisks indicate strains included in the susceptibility study. Clones are specified by a “c” followed by the clone number.
With a single exception, all strains included in susceptibility testing could be identified reliably to the species level by using the ITS region, because of their position in shared clades with ex-type strains (Fig. 2). The correct identification of Syncephalastrum racemosum turned out to be problematic because this species has not been typified yet, and strains morphologically identified as Syncephalastrum racemosum formed two clearly separate clades in the ITS tree (Fig. 2f): some S. racemosum strains were part of a well-supported clade with Syncephalastrum monosporum, while others (including the clinical strain CNRMA03.414), represented by two types of ITS sequences (note different clone numbers of the same strains), formed a second group. Syncephalastrum racemosum has been described by Schroeter (original reference cited in reference 13) as “inter Aspergillus oryzae in Oryza et pane,” i.e., on moldy rice and bread, in Wroclaw, Poland, and no type material is known to be preserved. None of the isolates studied morphologically completely matched the description given by Schroeter, and more-detailed taxonomic studies are needed to select a neotype for this species. Therefore, we designate the putative species in the following as Syncephalastrum “racemosum” species I and S. “racemosum” species II.
Judging from LSU and ITS phylograms (Fig. 1, 2) Rhizomucor regularior and Rhizomucor variabilis were found to be related to Mucor but belonged to the species complexes of M. circinelloides and M. hiemalis, respectively; R. variabilis has recently been renamed as M. irregularis, and R. regularior has been synonymized with M. circinelloides (9). The genus Rhizomucor is restricted to thermophilic species forming spherical spores, such as R. pusillus and R. miehei. Mucor ramosissimus belongs to M. circinelloides. The varieties azygosporus, chinensis, rhizopodiformis, and tuberosus of Rhizopus microsporus possessed identical ITS sequences and could not be discriminated molecularly (Fig. 2b). Therefore, the morphology-based assignment to the varieties was retained. In Rhizopus arrhizus (syn. R. oryzae), small molecular differences between varieties were observed, but because of the limited number of reference strains available representing each of the varieties, identifications were done at the species level only.
A total of eight antifungal compounds were tested on six mucoralean genera: Mucor and Rhizopus (Mucoraceae), Lichtheimia and Rhizomucor (Lichtheimiaceae), Syncephalastrum (Syncephalastraceae), and Cunninghamella (Cunninghamellaceae). The Cunninghamellaceae and the Syncephalastraceae are unigeneric, and conclusions drawn on the genus are also valid for the family. The positions of the genera in Tables 1 and 2 refer to their positions in the LSU phylogram of Fig. 1, except for Rhizomucor. Rhizomucor as a member of the Lichtheimiaceae is placed with Lichtheimia because its position closer to Circinella in the LSU phylogram is not supported. Within the genera, strains are arranged according to their position in the phylogenetic trees of Fig. 2. Because clinical breakpoints for filamentous fungi have not been assigned to the majority of antifungals, we refer to those given by Almyroudis et al. (5) and de Hoog et al. (17), viz. AMB ≤ 1; CAS ≤ 2; 5-FC ≤ 16; ITC ≤ 0.5; PCZ ≤ 0.5; and VRC ≤ 2.
Table 2.
MIC ranges and geometric means (GMs) for genera and species inferred from the phylogramsa
| Name (no. of strains) | MIC range (GM) (μg/ml) for: |
|||||||
|---|---|---|---|---|---|---|---|---|
| 5-FC | MCF | CAS | TBF | VCZ | ITZ | PCZ | AMB | |
| Mucor (12) | >64 (>64) | >4 (>4) | >4 (>4) | 0.125–32 (2.69) | 1–>8 (>6.0) | 0.25–4 (1.19) | 0.125–2 (0.79) | 0.25–0.5 (0.37) |
| Mucor circinelloides sensu lato (8) | >64 (>64) | >4 (>4) | >4 (>4) | 2–32 (8) | >8 (>8) | 1–4 (1.68) | 0.5–2 (1) | 0.25–0.5 (0.32) |
| Mucor racemosus (1) | >64 | >4 | >4 | 0.125 | 2 | 0.25 | 0.125 | 0.5 |
| Mucor hiemalis (1) | >64 | >4 | >4 | 0.25 | 1 | 0.5 | 0.5 | 0.5 |
| Mucor irregularis (1) | >64 | >4 | >4 | 0.5 | >8 | 1 | 1 | 0.5 |
| Mucor indicus (1) | >64 | >4 | >4 | 0.5 | >8 | 1 | 1 | 0.5 |
| Rhizopus (19) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–32 (3.84) | 4–>8 (>5.5) | 0.125–4 (0.58) | 0.03–2 (0.33) | 0.5–2 (1) |
| Rhizopus arrhizus (14) | >64 (>64) | >4 (>4) | >4 (>4) | 1–32 (21.11) | 4–>8 (>5.9) | 0.125–1 (0.5) | 0.03–0.5 (0.28) | 0.5–2 (0.95) |
| Rhizopus microsporus (5) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.25 (0.12) | 4–8 (4.6) | 0.5–4 (0.87) | 0.25–2 (0.57) | 1–2 (1.15) |
| Syncephalastrum (6) | ≥64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.25 (0.18) | 1–>8 (>5.6) | 0.03–0.5 (0.28) | 0.06–1 (0.31) | 0.03–0.25 (0.09) |
| Syncephalastrum “racemosum” species II (2) | ≥64 | >4 | >4 | 0.25 | ≥8 | 0.5 | 0.25–5 | 0.03–0.06 |
| Syncephalastrum “racemosum” species I (4) | ≥64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.25 (0.15) | 1–>8 (>4.7) | 0.03–0.5 (0.21) | 0.06–1 (0.29) | 0.06–0.25 (0.12) |
| Rhizomucor (8) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.25 (0.11) | 2–>8 (>3.1) | 0.06–0.25 (0.21) | 0.125–0.5 (0.21) | 0.5 (0.5) |
| Rhizomucor pusillus (7) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.125 (0.1) | 2–>8 (>3.0) | 0.25 (0.25) | 0.125–0.5 (0.23) | 0.5 (0.5) |
| Rhizomucor miehei (1) | >64 | >4 | >4 | 0.25 | 4 | 0.06 | 0.125 | 0.5 |
| Lichtheimia (13) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.5 (0.18) | 2–>8 (>5.8) | 0.03–0.5 (0.18) | 0.06–0.25 (0.13) | 0.06–0.5 (0.24) |
| Lichtheimia corymbifera (6) | >64 (>64) | >4 (>4) | >4 (>4) | 0.06–0.25 (0.12) | ≥8 (>8) | 0.06–0.5 (0.22) | 0.06–0.25 (0.20) | 0.25–0.5 (0.31) |
| Lichtheimia ramosa (5) | >64 (>64) | >4 (>4) | >4 (>4) | 0.125–0.5 (0.25) | 2–>8 (>4.6) | 0.03–0.5 (0.19) | 0.06–0.25 (0.11) | 0.06–0.25 (0.12) |
| Lichtheimia ornata (2) | >64 | >4 | >4 | 0.25 | 2–>8 | 0.06–0.125 | 0.06 | 0.25 |
| Cunninghamella (8) | 32–>64 (>58.7) | 0.5–>4 (>2.59) | ≥4 (>4) | 0.06–0.25 (0.09) | >8 (>8) | 1–>64 (>19.0) | 0.5–1 (0.71) | 1–2 (1.54) |
| Cunninghamella bertholletiae (5) | 32–>64 (>55.7) | 1–>4 (>3.03) | ≥4 (>4) | 0.06–0.25 (0.08) | >8 (>8) | 16–>64 (>27.9) | 0.5–1 (0.66) | 1–2 (1.52) |
| Cunninghamella echinulata (3) | >64 | 0.5–>4 | ≥4 | 0.06–0.3 | >8 | 1–>64 | 0.5–1 | 1–2 |
| All isolates (66) | 32–>64 (>63.3) | 0.5–>4 (>3.8) | ≥4 (>4) | 0.06–32 (0.62) | 1–>8 (>5.5) | 0.03–>64 (>0.66) | 0.03–2 (0.33) | 0.03–2 (0.48) |
MIC ranges (GMs) for 5-fluorocytosine (5-FC), micafungin (MCF), caspofungin (CAS), terbinafine (TBF), voriconazole (VCZ), itraconazole (ITZ), posaconazole (PCZ), and amphotericin B (AMB). Genera are placed in line with the phylogenetic tree of Fig. 1 except for Rhizomucor. Within the genera, species are arranged according to their position in the trees of Fig. 2.
For 5-FC, MCF, and CAS, little variation in susceptibility was noted: for 5-FC and CAS, all strains showed high MIC/MEC values of 32 to >64 and ≥4 μg/ml, respectively. Except for two strains of Cunninghamella, high MEC values (≥4 μg/ml) were also obtained for MCF. All strains included in this study were resistant or less susceptible to VCZ.
For the remaining compounds, considerable variation was found at the family and generic levels inferred from the LSU phylogram. Overall, AMB in vitro was the most effective antifungal agent against Mucorales, but its efficacy proved to be ambiguous in Rhizopus and Cunninghamella, where relatively high MICs were obtained. Three out of 19 Rhizopus strains and 5 out of 8 Cunninghamella strains tested had MICs of 2 μg/ml, exceeding the assumed breakpoint for this drug. MICs of these two genera were significantly elevated when compared to the other Mucorales, for which the amphotericin B MICs ranged between 0.03 and 0.5 μg/ml (Mann-Whitney, P < 0.0001 for both genera). Together with those for posaconazole, amphotericin B MICs showed the smallest range of variation over the entire set of Mucorales tested (Table 2). Among the azoles, PCZ was the most effective antifungal drug, with all strains being inhibited by concentrations of 2 μg/ml. The genera Mucor (defined in a phylogenetic sense) and Cunninghamella showed significantly lower degrees of susceptibility to PCZ (Mann-Whitney, P < 0.0001 for both genera). In Rhizopus and Syncephalastrum, resistant strains occurred only sporadically. The Lichtheimiaceae, represented by Lichtheimia and Rhizomucor, completely lacked strains with reduced susceptibility for PCZ.
Itraconazole was significantly less active against the Mucoraceae (defined in a phylogenetic sense) and Cunninghamella, in which especially high MIC values were obtained (Mann-Whitney; P < 0.0001 for the Mucoraceae as well as for Cunninghamella). In contrast, the Lichtheimiaceae and Syncephalastrum (Syncephalastraceae), united in a well-supported clade in the LSU phylogram, did not include strains with reduced susceptibility for ITZ.
For TBF, elevated MIC values were found only in the Mucoraceae, while the remaining families, namely, the Cunninghamellaceae, the Lichtheimiaceae, and the Syncephalastraceae, were fully susceptible. The susceptibility of Cunninghamella to TBF is of special importance because TBF was the only thoroughly active antifungal against this genus in our study. The susceptibility to TBF in the Mucoraceae appeared to be species dependent. Terbinafine MICs were strikingly different between R. microsporus (geometric mean [GM] MIC of 0.12 μg/ml) and R. arrhizus (GM MIC of 21.1 μg/ml) (Mann-Whitney, P = 0.0020) but not between M. circinelloides sensu lato (GM MIC of 5.66 μg/ml) and the remaining Mucor species (MICs of ≤0.5 μg/ml), although a trend was noted (Mann-Whitney, P = 0.0746). Susceptibility values of PCZ and TBF were more heterogeneous in the Mucoraceae than in the remaining families.
Maximum susceptibilities were reached with PCZ (MIC = 0.03 μg/ml) for Rhizopus arrhizus, ITZ (MIC = 0.03 μg/ml) for Lichtheimia ramosa and Syncephalastrum “racemosum” species I, and AMB (MIC = 0.03 μg/ml) for Syncephalastrum “racemosum” species II, while maximum resistance was observed with 5-FC (MIC > 64 μg/ml) for nearly all strains tested and with ITZ (MIC > 64 μg/ml) for 3 Cunninghamella strains.
Intrageneric differences of in vitro susceptibility in Mucor and Rhizopus ranged from 4 to 9 log2 dilution steps. With respect to PCZ and ITZ, the genus Mucor seems to contain a higher proportion of strains that are resistant to or show reduced susceptibility than Rhizopus (Mann-Whitney, P = 0.0042 and P = 0.0369, respectively). In contrast MICs for VCZ were lower in Mucor than in Rhizopus (Mann-Whitney, P < 0.0001). Some intraspecific variability of in vitro susceptibility was found in all genera tested (Table 1). Strain CNRMA 04.1469 of R. microsporus deviated from remaining strains of this species in its response to azoles.
DISCUSSION
A robust taxonomy and phylogeny of the Mucorales require species validation by their ex-type materials. These reference materials serve to calibrate additional strains examined, such that a taxonomic scaffold is provided, allowing meaningful comparison of susceptibility data at the levels of species, genera, and families.
The genera Mucor and Rhizomucor have been misapplied in some medical publications. Judging from ITS sequence data, Rhizomucor regularior (as R. variabilis var. regularior) was declared to be a synonym of Mucor circinelloides (9) as suggested before (6, 36). The typical variety of Rhizomucor variabilis var. variabilis fits elsewhere in the genus Mucor (1, 10, 42), and recently Álvarez et al. (9) proposed a name change to Mucor irregularis.
Our study supports AMB as the antifungal of choice for most genera of Mucorales. High MICs for AMB have been reported for some genera not tested in the present study, such as Apophysomyces and Saksenaea (7, 8, 12, 15, 35, 39). Results for Rhizopus and Cunninghamella, however, are ambiguous, with MICs of 2 μg/ml for AMB. Similar results were obtained by other authors (1, 35, 39, 40). Alastruey-Izquierdo et al. (1) listed R. arrhizus and Cunninghamella bertholletiae with MIC ranges as wide as 0.03 to 32 μg/ml and 2 to 32 μg/ml, respectively. In concordance with previous studies (1, 5, 16), PCZ was the second most active drug, all strains being inhibited by 2 μg/ml or less (Table 2). The success rate of therapy was reported to be 79% (20).
5-FC, MCF, and CAS were ineffective in all or nearly all Mucorales, showing high MIC/MEC values of ≥32, ≥4 (for 64 out of 66 strains), and ≥4 μg/ml, respectively. This resistance is in accordance with literature data (1, 5, 16, 40). With MICs consistently ≥1 μg/ml in all Mucorales analyzed, VCZ was less active than other azoles. The poor activity of this antifungal has been highlighted previously (e.g., references 1, 5, and 16).
Phylogenetic distances within the Mucorales are reflected in in vitro susceptibility profiles against antifungal drugs on all levels of family, genus, species, and strain (Tables 1 and 2). Members of the Lichtheimiaceae were consistently susceptible to PCZ, ITZ, TBF, and AMB and exhibited smaller MIC ranges than the Mucoraceae. This matches with MIC values published (5, 16, 24, 32, 35, 39), although Torres-Narbona et al. (40) and Alastruey-Izquierdo et al. (1, 4) reported on individual Lichtheimia strains that were resistant to PCZ, ITZ, and TBF and to ITZ, respectively.
The consistent in vitro activity of TBF against Cunninghamella is of clinical relevance because the proportion of strains that are resistant or have reduced susceptibility to PCZ and AMB is relatively high in this genus, as shown in this study and by other authors (1, 5, 39). Alastruey-Izquierdo et al. (1) tested the susceptibility of Cunninghamella against TBF with a similar result. However, the authors found an individual strain resistant to TBF.
The species-specific differences in the susceptibility to TBF found (Rhizopus arrhizus [resistant] versus R. microsporus [susceptible] and Mucor circinelloides sensu lato [resistant] versus remaining Mucor species [susceptible]) were also in agreement with earlier studies (1, 16). Significantly deviating resistance to azoles was found in a single strain of R. microsporus; Alastruey-Izquierdo et al. (1) reported that TBF data were variable in that species.
In conclusion, AMB and PCZ were the most effective antifungal agents against Mucorales. Susceptibility profiles (restricted to the drugs that were at least partly active) differed significantly at the familial, generic, and specific levels and reflected relationships as referred from the phylogram; the Lichtheimiaceae were fully susceptible to PCZ, ITZ, TBF, and AMB. Syncephalastrum (Syncephalastraceae) members, positioned at the shortest distance to the Lichtheimiaceae in the LSU phylogram, showed similar susceptibility profiles. The only difference was a single strain resistant to PCZ. In contrast, the Mucoraceae were characterized by a reduced susceptibility to PCZ, ITZ, and AMB and a lack of activity of TBF in some species. While more Mucor strains were resistant to the azoles, only Rhizopus strains exhibited a reduced susceptibility to AMB. Susceptibility profiles of Cunninghamella (Cunninghamellaceae) resembled those of the Mucoraceae in terms of the high proportion of strains resistant to PCZ, ITZ, and AMB but differed in terms of the low MIC values for TBF. Judging from MIC values recorded by other authors, the Sakseneaeceae behave similarly to Cunninghamella, their closest studied neighbor in the LSU phylogram, in susceptibility tests. Along with Cunninghamella, they exhibit a lack of or reduced susceptibility against PCZ, ITZ, and AMB (5, 7, 8, 12, 35) and low MIC values for TBF (7) (tested only for Saksenaea).
Obviously, the Mucorales cannot be considered as a single entity from an antifungal perspective, since large differences in susceptibility exist between families, genera, species, and strains. The reduced activity of AMB in Rhizopus and Cunninghamella and the high proportion of Mucor and Cunninghamella strains that were less susceptible to PSZ are of practical importance because they concern deviations for compounds that are otherwise recommended for antifungal therapy of infections due to Mucorales. Molecular identification of the etiologic agent is therefore required unless the susceptibility profile of the strain is known. In genera with high intraspecific variation in their antifungal profiles, such as Mucor and Rhizopus, immediate susceptibility testing is recommended to confirm the most effective and appropriate compound.
ACKNOWLEDGMENTS
We are indebted to Ana Alastruey-Izquierdo and Xuelian Lu for useful comments on the manuscript.
J.F.M. has been a consultant to Astellas, Basilea, Merck, and Schering-Plough and received speaker's fees from Gilead, Janssen Pharmaceutica, Merck, Pfizer, and Schering-Plough.
Footnotes
Published ahead of print 9 November 2011
REFERENCES
- 1. Alastruey-Izquierdo A, et al. 2009. Activity of posaconazole and other antifungal agents against Mucorales strains identified by sequencing of internal transcribed spacers. Antimicrob. Agents Chemother. 53:1686–1689 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2. Alastruey-Izquierdo A, et al. 2009. In vitro activity of antifungals against zygomycetes. Clin. Microbiol. Infect. 15(Suppl. 5):71–76 [DOI] [PubMed] [Google Scholar]
- 3. Alastruey-Izquierdo A, et al. 2010. Species recognition and clinical relevance of the zygomycetous genus Lichtheimia (syn, Absidia pro parte, Mycocladus). J. Clin. Microbiol. 48:2154–2170 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4. Alastruey-Izquierdo A, Cuesta I, Walther G, Cuenca-Estrella M, Rodriguez-Tudela JL. 2010. Antifungal susceptibility profile of human-pathogenic species of Lichtheimia. Antimicrob. Agents Chemother. 54:3058–3060 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 5. Almyroudis NG, Sutton DA, Fothergill AW, Rinaldi MG, Kusne S. 2007. In vitro susceptibilities of 217 clinical isolates of zygomycetes to conventional and new antifungal agents. Antimicrob. Agents Chemother. 51:2587–2590 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6. Álvarez E, et al. 2009. Spectrum of zygomycete species identified from clinically significant specimens in the United States. J. Clin. Microbiol. 47:1650–1656 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7. Álvarez E, et al. 2010. Molecular phylogeny and proposal of two new species of the emerging pathogenic fungus Saksenaea. J. Clin. Microbiol. 48:4410–4416 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8. Álvarez E, et al. 2010. Molecular phylogenetic diversity of the emerging mucoralean fungus Apophysomyces: proposal of three new species. Rev. Iberoam. Micol. 27:80–89 [DOI] [PubMed] [Google Scholar]
- 9. Álvarez E, et al. 2011. Two new species of Mucor from clinical samples. Med. Mycol. 49:62–72 [DOI] [PubMed] [Google Scholar]
- 10. Boekhout T, et al. 2009. Fungal taxonomy: new developments in medically important fungi. Curr. Fung. Infect. Rep. 3:170–178 [Google Scholar]
- 11. Boelaert JR, et al. 1993. Mucormycosis during deferoxamine therapy is a siderophore-mediated infection. In vitro and in vivo animal studies. J. Clin. Invest. 91:1979–1986 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12. Chakrabarti A, et al. 2010. Apophysomyces elegans: epidemiology, AFLP typing, and in vitro antifungal susceptibility pattern. J. Clin. Microbiol. 48:4580–4585 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13. Cohn FJ. (ed). 1886. Kryptogamen-Flora von Schlesien, vol 3, first half. Kern's Verlag, Breslau, Germany [Google Scholar]
- 14. Däbritz J, et al. 2011. Mucormycosis in paediatric patients: demographics, risk factors and outcome of 12 contemporary cases. Mycoses 54:e785–e788 [DOI] [PubMed] [Google Scholar]
- 15. Dannaoui E, Meis JF, Mouton JW, Verweij PE. 2002. In vitro susceptibilities of Zygomycota to polyenes. J. Antimicrob. Chemother. 49:741–744 [DOI] [PubMed] [Google Scholar]
- 16. Dannaoui E, Meletiadis J, Mouton JW, Meis JF, Verweij PE. 2003. In vitro susceptibilities of zygomycetes to conventional and new antifungals. J. Antimicrob. Chemother. 51:45–52 [DOI] [PubMed] [Google Scholar]
- 17. de Hoog GS, Guarro J, Gené J, Figueras MJ. 2000. Atlas of clinical fungi, 2nd ed. Reus, Utrecht, The Netherlands [Google Scholar]
- 18. de Hoog GS, Gerrits van den Ende AHG. 1998. Molecular diagnostics of clinical strains of filamentous basidiomycetes. Mycoses 41:183–189 [DOI] [PubMed] [Google Scholar]
- 19. Goel S, Carter JE, Culpepper M, Kahn AG. 2009. Primary renal zygomycotic infarction mimicking renal neoplasia in an immunocompetent patient. Am. J. Med. Sci. 338:330–333 [DOI] [PubMed] [Google Scholar]
- 20. Greenberg RN, et al. 2006. Posaconazole as salvage therapy for zygomycosis. Antimicrob. Agents Chemother. 50:126–190 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21. Gupta KL, et al. 1999. Renal zygomycosis: an under-diagnosed cause of acute renal failure. Nephrol. Dial. Transpl. 14:2720–2725 [DOI] [PubMed] [Google Scholar]
- 22. Hata DJ, Buckwalter SP, Pritt BS, Roberts GD, Wengenack NL. 2008. Real-time PCR method for detection of zygomycetes. J. Clin. Microbiol. 46:2353–2358 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 23. Hibbett DS, et al. 2007. A higher-level phylogenetic classification of the Fungi. Mycol. Res. 111:509–547 [DOI] [PubMed] [Google Scholar]
- 24. Johnson EM, Szekely A, Warnock DW. 1998. In-vitro activity of voriconazole, itraconazole and amphotericin B against filamentous fungi. J. Antimicrob. Chemother. 42:741–745 [DOI] [PubMed] [Google Scholar]
- 25. Lu XL, et al. 2009. Primary cutaneous zygomycosis caused by Rhizomucor variabilis: a new endemic zygomycosis? A case report and review of 6 cases reported from China. Clin. Infect. Dis. 49:e39–e43 [DOI] [PubMed] [Google Scholar]
- 26. Maddox L, Long GD, Vredenburgh JJ, Folz RJ. 2001. Rhizopus presenting as an endobronchial obstruction following bone marrow transplant. Bone Marrow Transpl. 28:634–636 [DOI] [PubMed] [Google Scholar]
- 27. Marak RSK, et al. 2010. Successful medical management of renal zygomycosis: a summary of two cases and a review of the Indian literature. Med. Mycol. 48:1088–1095 [DOI] [PubMed] [Google Scholar]
- 28. Möller EM, Bahnweg G, Sandermann H, Geiger HH. 1992. A simple and efficient protocol for isolation of high molecular weight DNA from filamentous fungi, fruit bodies, and infected plant tissues. Nucleic Acids Res. 22:6115–6116 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29. NCCLS/CLSI 2005. Reference method for broth dilution antifungal susceptibility. Testing of filamentous fungi. Approved standard M38-A. Clinical and Laboratory Standards Institute, Wayne, PA [Google Scholar]
- 30. O'Donnell K. 1993. Fusarium and its near relatives, p 225–233 In Reynolds DR, Taylor JW. (ed), The fungal holomorph: mitotic, meiotic and pleomorphic speciation in fungal systematics. CAB International, Wallingford, United Kingdom [Google Scholar]
- 31. O'Donnell K, Lutzoni FM, Ward TJ, Benny GL. 2001. Evolutionary relationships among mucoralean fungi (Zygomycota): evidence for family polyphyly on a large scale. Mycologia 93:286–296 [Google Scholar]
- 32. Otcenasek M, Buchta V. 1994. In vitro susceptibility to 9 antifungal agents of 14 strains of zygomycetes isolated from clinical specimens. Mycopathologia 128:135–137 [DOI] [PubMed] [Google Scholar]
- 33. Rambaut A. 2002. Se-Al. http://tree.bio.ed.ac.uk/software/seal/
- 34. Roden MM, et al. 2005. Epidemiology and outcome of zygomycosis: a review of 929 reported cases. Clin. Infect. Dis. 41:634–653 [DOI] [PubMed] [Google Scholar]
- 35. Sabatelli F, et al. 2006. In vitro activities of posaconazole, fluconazole, itraconazole, voriconazole, and amphotericin B against a large collection of clinically important molds and yeasts. Antimicrob. Agents Chemother. 50:2009–2015 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 36. Schwarz P, et al. 2006. Molecular identification of zygomycetes from culture and experimentally infected tissue. J. Clin. Microbiol. 44:340–349 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 37. Skiada A, et al. 2011. Zygomycosis in Europe: analysis of 230 cases accrued by the registry of the European Confederation of Medical Mycology (ECMM) Working Group on Zygomycosis between 2005 and 2007. Clin. Microbiol. Infect. 17:1859–1867 [DOI] [PubMed] [Google Scholar]
- 38. Stamatakis A, Hoover P, Rougemont J. 2008. A rapid bootstrap algorithm for the RAxML Web-Servers. Syst. Biol. 75:758–771 [DOI] [PubMed] [Google Scholar]
- 39. Sun QN, Fothergill AW, McCarthy DI, Rinaldi MG, Graybill JR. 2002. In vitro activities of posaconazole, itraconazole, voriconazole, amphotericin B, and fluconazole against 37 clinical isolates of zygomycetes. Antimicrob. Agents Chemother. 46:1581–1582 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 40. Torres-Narbona M, Guinea J, Martinez-Alarcon J, Pelaez T, Bouza E. 2007. In vitro activities of amphotericin B, caspofungin, itraconazole, posaconazole, and voriconazole against 45 clinical isolates of zygomycetes: comparison of CLSI M38-A, Sensititre YeastOne, and the Etest. Antimicrob. Agents Chemother. 51:1126–1129 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 41. Vilgalys R, Hester M. 1990. Rapid genetic identification and mapping of enzymatically amplified ribosomal DNA from several Cryptococcus species. J. Bacteriol. 172:4238–4246 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 42. Voigt K, Cigelnik E, O'Donnell K. 1999. Phylogeny and PCR identification of clinically important zygomycetes based on nuclear ribosomal-DNA sequence data. J. Clin. Microbiol. 37:3957–3964 [DOI] [PMC free article] [PubMed] [Google Scholar]
- 43. Voigt K, Wöstemeyer J. 2001. Phylogeny and origin of 82 zygomycetes from all 54 genera of the Mucorales and Mortierellales based on combined analysis of actin and translation elongation factor EF-1a genes. Gene 270:113–120 [DOI] [PubMed] [Google Scholar]
- 44. White TJ, Bruns T, Lee S, Taylor JW. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics, pp. 315–322 In Innis MA, Gelfand DH, Sninsky JJ, White TJ. (ed), PCR protocols: a guide to methods and applications. Academic Press, Inc., New York, NY [Google Scholar]