Use of Commercially Available Cryogenic Vials for
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微生物临床杂志 2006年第2期
ABSTRACT
The use of commercially available cryogenic vials (Microbank vials) stored at –70°C for the storage and preservation of dermatophyte fungi was investigated. None of the 200 strains of dermatophytes examined, representing 21 species, showed a loss of viability after they had been stored for periods ranging from 1 week to 2 years at –70°C. All strains showed typical colonial and microscopic morphologies following revival.
TEXT
Long-term storage of fungal isolates is critical for preservation of the germplasm and maintenance of stock cultures with minimal effort over long periods. Conservation of morphological, physiological, genetic, and metabolic stability is crucial for many purposes and is vital for isolates used as medical reference strains, for chemotaxonomic studies, or in the commercial production of biochemicals. In a comparative study (3) of the effects of five different storage methods, cryopreservation was the method that best provided for the stability of secondary metabolite production and is now considered the best method available for the long-term storage of microbial cultures (4). Dermatophytic fungi can present problems for storage, as the cultures often become pleomorphic, with various levels of sporulation or mycelial growth. McGinnis et al. (2) used sterile distilled water to store hyphal and spore suspensions of 147 different species of fungi at 25°C for periods of 12 to 60 months. These included more than 25 species of dermatophytes. The degree of sporulation and the quality of the inoculum appeared to be critical factors; and when the inocula were "adequate" in size, even some poorly sporulating species such as Trichophyton violaceum, Trichophyton schoenleinii, and Microsporum ferrugineum survived storage well. The long-term storage of a wide range of fungal species, including dermatophyte isolates, in commercially prepared cryogenic freezer beads (Microbank) at –70°C or in liquid nitrogen has been tested (1). Although most fungi were preserved well by this method, dermatophytic fungi did not show good recovery rates. For example, more than 50% of isolates of Trichophyton rubrum were not recovered. Consequently, this method was not recommended for use for the long-term storage of dermatophytic fungi. In contrast, we have found contradictory results and we report on a simple and successful technique for the long-term storage of dermatophytes.
Fresh isolates were collected from clinical specimens submitted to the Microbiology Department at the William Harvey Hospital, Ashford, United Kingdom, between March 2002 and August 2005. Reference strains were obtained from the National Collection of Pathogenic Fungi, Bristol, United Kingdom. Representative strains of dermatophytes were used to assess a commercially available freezer bead storage kit (Microbank; Pro-Labs Diagnostics, Richmond Hill, Ontario, Canada). Each 2-ml tube contains approximately 50 plastic beads (diameter, 3 mm) with a hole through the center (this hole retains approximately 1 μl of suspension), which allows repeated recovery of an isolate before the preparation of a new stored culture is needed. This is in contrast to traditional long-term storage methods, in which the isolates are stored in multiple single-use vials, and has the added advantage of taking up less space.
Mycelium and conidia were harvested from 7-day-old cultures incubated at 27°C on Sabouraud dextrose agar (SDA; Oxoid Ltd., United Kingdom) by using a sterile scalpel and inoculated into a freezer bead tube containing a suspension medium prepared according to the manufacturer's instructions to give a density approximately equal to or greater than that of a McFarland no. 4 standard. The suspension was shaken vigorously to evenly distribute the fungus and was left to stand for 5 min. Excess fluid was removed with a Pasteur pipette. Before the tubes were frozen and stored at –70°C, a single bead was removed with sterile forceps and was placed on a fresh SDA plate, and the resulting drop of fungal suspension was spread by using a 10-μl loop to obtain single colonies and to check for viability and purity. These plates were incubated at 27°C for 7 days to assess the amount of inoculum present on a single bead. At various time intervals over 24 months, the tubes were removed from the freezer and a bead was removed from the frozen clump, plated, and incubated as described above. The tubes were immediately returned to the –70°C freezer before the contents had thawed. The number of colonies recovered, their growth rate, and the macroscopic and microscopic morphologies of the isolates were noted.
A detailed time course study was conducted with four isolates (Trichophyton interdigitale WHH1268, Trichophyton mentagrophytes WHH692, T. rubrum WHH3229, and Epidermophyton floccosum WHH1471). Single beads were removed at 0, 1, 2, 3, 4, 5, 6, 7, 8, 12, 16, 20, 24, 32, and 36 weeks and were cultured as described above. Epidermophyton floccosum was included, as it is known to die rapidly if it is kept at 4°C. In all cases, at all time intervals, successful reestablishment of the cultures ensued. At least 1,000 CFU was typically recovered from each bead. The growth rates and the hyphal densities were comparable to those of an initial control culture before it was frozen. Colonial and microscopic morphologies remained true to type throughout.
In addition to this time course study, a second trial was conducted with 58 stored isolates representing 15 species of dermatophytes. The isolates were selected to give a range of species but also to sample a range of isolates within some of these species (e.g., Arthroderma benhamiae, Trichophyton interdigitale, and Trichophyton tonsurans). For this trial, isolates stored for different time periods over the previous 24 months were recultured in triplicate to assess the uniformity of the distribution of viable organisms in frozen tubes. Three freezer beads were taken from each tube of preserved isolates and cultured as described above. In all cases, successful reestablishment from all three replicates occurred for all isolates tested. No adverse effects on morphology or growth rates compared to those of cultures not subjected to cryopreservation were noted.
Following these initial trials, this method of preservation was adopted for the storage of all stock strains in the laboratory. To date, all cultures kept in this manner have been successfully revived as required at times ranging from 1 week to 2 years, and these cultures represent 200 isolates of 21 species of dermatophytes (Table 1). Espinel-Ingroff et al. (1) have discussed the advantages of using the Microbank freezer bead system in terms of its availability and ease of use. Their results suggested that dermatophytes would not be well preserved by use of this method. Our results for a wider range of isolates suggest that the limited numbers of dermatophytes that they tested were not representative or that a different preservation technique might have given better recovery rates. For example, half the specimens prepared by Epinel-Ingroff et al. (1) were preserved in liquid nitrogen and kept for up to 8 years, whereas all our specimens were kept at –70°C and tested within 2 years. The recovery of all the isolates used in our study was successful, with no apparent effect on culture phenotype. All four specimens of T. rubrum were recovered in our study, including the isolate used in the detailed time course study. This is in contrast to the 54% recovery rate of T. rubrum isolates in the earlier study (1). As a consequence, we can now recommend this method of preservation of dermatophytes for clinical laboratories worldwide.
ACKNOWLEDGMENTS
We acknowledge the help of the East Kent NHS Trust in providing funds to support this work.
REFERENCES
Espinel-Ingroff, A., D. Montero, and E. Marti-Mazuelos. 2004. Long-term preservation of fungal isolates in commercially prepared cryogenic Microbank vials. J. Clin. Microbiol. 42:1257-1259.
McGinnis, M. R., A. A. Padhye, and L. Ajello. 1974. Storage of stock cultures of filamentous fungi, yeasts and some aerobic actinomycetes in sterile distilled water. Appl. Microbiol. 28:218-222.
Ryan, M. J., P. D. Bridge, D. Smith, and P. Jeffries. 2002. Phenotypic degeneration occurs during sector formation in Metarhizium anisopliae. J. Appl. Microbiol. 93:163-168.
Ryan, M. J., P. Jeffries, P. D. Bridge, and D. Smith. 2001. Developing cryopreservation protocols to secure fungal gene function. Cryo Lett. 22:115-124.
East Kent Microbiology Service, The William Harvey Hospital, Kennington Road, Ashford, Kent TN24 0LZ
1 Department of Biosciences, University of Kent, Canterbury, Kent CT2 6NJ, United Kingdom(M. Baker and P. Jeffries)
The use of commercially available cryogenic vials (Microbank vials) stored at –70°C for the storage and preservation of dermatophyte fungi was investigated. None of the 200 strains of dermatophytes examined, representing 21 species, showed a loss of viability after they had been stored for periods ranging from 1 week to 2 years at –70°C. All strains showed typical colonial and microscopic morphologies following revival.
TEXT
Long-term storage of fungal isolates is critical for preservation of the germplasm and maintenance of stock cultures with minimal effort over long periods. Conservation of morphological, physiological, genetic, and metabolic stability is crucial for many purposes and is vital for isolates used as medical reference strains, for chemotaxonomic studies, or in the commercial production of biochemicals. In a comparative study (3) of the effects of five different storage methods, cryopreservation was the method that best provided for the stability of secondary metabolite production and is now considered the best method available for the long-term storage of microbial cultures (4). Dermatophytic fungi can present problems for storage, as the cultures often become pleomorphic, with various levels of sporulation or mycelial growth. McGinnis et al. (2) used sterile distilled water to store hyphal and spore suspensions of 147 different species of fungi at 25°C for periods of 12 to 60 months. These included more than 25 species of dermatophytes. The degree of sporulation and the quality of the inoculum appeared to be critical factors; and when the inocula were "adequate" in size, even some poorly sporulating species such as Trichophyton violaceum, Trichophyton schoenleinii, and Microsporum ferrugineum survived storage well. The long-term storage of a wide range of fungal species, including dermatophyte isolates, in commercially prepared cryogenic freezer beads (Microbank) at –70°C or in liquid nitrogen has been tested (1). Although most fungi were preserved well by this method, dermatophytic fungi did not show good recovery rates. For example, more than 50% of isolates of Trichophyton rubrum were not recovered. Consequently, this method was not recommended for use for the long-term storage of dermatophytic fungi. In contrast, we have found contradictory results and we report on a simple and successful technique for the long-term storage of dermatophytes.
Fresh isolates were collected from clinical specimens submitted to the Microbiology Department at the William Harvey Hospital, Ashford, United Kingdom, between March 2002 and August 2005. Reference strains were obtained from the National Collection of Pathogenic Fungi, Bristol, United Kingdom. Representative strains of dermatophytes were used to assess a commercially available freezer bead storage kit (Microbank; Pro-Labs Diagnostics, Richmond Hill, Ontario, Canada). Each 2-ml tube contains approximately 50 plastic beads (diameter, 3 mm) with a hole through the center (this hole retains approximately 1 μl of suspension), which allows repeated recovery of an isolate before the preparation of a new stored culture is needed. This is in contrast to traditional long-term storage methods, in which the isolates are stored in multiple single-use vials, and has the added advantage of taking up less space.
Mycelium and conidia were harvested from 7-day-old cultures incubated at 27°C on Sabouraud dextrose agar (SDA; Oxoid Ltd., United Kingdom) by using a sterile scalpel and inoculated into a freezer bead tube containing a suspension medium prepared according to the manufacturer's instructions to give a density approximately equal to or greater than that of a McFarland no. 4 standard. The suspension was shaken vigorously to evenly distribute the fungus and was left to stand for 5 min. Excess fluid was removed with a Pasteur pipette. Before the tubes were frozen and stored at –70°C, a single bead was removed with sterile forceps and was placed on a fresh SDA plate, and the resulting drop of fungal suspension was spread by using a 10-μl loop to obtain single colonies and to check for viability and purity. These plates were incubated at 27°C for 7 days to assess the amount of inoculum present on a single bead. At various time intervals over 24 months, the tubes were removed from the freezer and a bead was removed from the frozen clump, plated, and incubated as described above. The tubes were immediately returned to the –70°C freezer before the contents had thawed. The number of colonies recovered, their growth rate, and the macroscopic and microscopic morphologies of the isolates were noted.
A detailed time course study was conducted with four isolates (Trichophyton interdigitale WHH1268, Trichophyton mentagrophytes WHH692, T. rubrum WHH3229, and Epidermophyton floccosum WHH1471). Single beads were removed at 0, 1, 2, 3, 4, 5, 6, 7, 8, 12, 16, 20, 24, 32, and 36 weeks and were cultured as described above. Epidermophyton floccosum was included, as it is known to die rapidly if it is kept at 4°C. In all cases, at all time intervals, successful reestablishment of the cultures ensued. At least 1,000 CFU was typically recovered from each bead. The growth rates and the hyphal densities were comparable to those of an initial control culture before it was frozen. Colonial and microscopic morphologies remained true to type throughout.
In addition to this time course study, a second trial was conducted with 58 stored isolates representing 15 species of dermatophytes. The isolates were selected to give a range of species but also to sample a range of isolates within some of these species (e.g., Arthroderma benhamiae, Trichophyton interdigitale, and Trichophyton tonsurans). For this trial, isolates stored for different time periods over the previous 24 months were recultured in triplicate to assess the uniformity of the distribution of viable organisms in frozen tubes. Three freezer beads were taken from each tube of preserved isolates and cultured as described above. In all cases, successful reestablishment from all three replicates occurred for all isolates tested. No adverse effects on morphology or growth rates compared to those of cultures not subjected to cryopreservation were noted.
Following these initial trials, this method of preservation was adopted for the storage of all stock strains in the laboratory. To date, all cultures kept in this manner have been successfully revived as required at times ranging from 1 week to 2 years, and these cultures represent 200 isolates of 21 species of dermatophytes (Table 1). Espinel-Ingroff et al. (1) have discussed the advantages of using the Microbank freezer bead system in terms of its availability and ease of use. Their results suggested that dermatophytes would not be well preserved by use of this method. Our results for a wider range of isolates suggest that the limited numbers of dermatophytes that they tested were not representative or that a different preservation technique might have given better recovery rates. For example, half the specimens prepared by Epinel-Ingroff et al. (1) were preserved in liquid nitrogen and kept for up to 8 years, whereas all our specimens were kept at –70°C and tested within 2 years. The recovery of all the isolates used in our study was successful, with no apparent effect on culture phenotype. All four specimens of T. rubrum were recovered in our study, including the isolate used in the detailed time course study. This is in contrast to the 54% recovery rate of T. rubrum isolates in the earlier study (1). As a consequence, we can now recommend this method of preservation of dermatophytes for clinical laboratories worldwide.
ACKNOWLEDGMENTS
We acknowledge the help of the East Kent NHS Trust in providing funds to support this work.
REFERENCES
Espinel-Ingroff, A., D. Montero, and E. Marti-Mazuelos. 2004. Long-term preservation of fungal isolates in commercially prepared cryogenic Microbank vials. J. Clin. Microbiol. 42:1257-1259.
McGinnis, M. R., A. A. Padhye, and L. Ajello. 1974. Storage of stock cultures of filamentous fungi, yeasts and some aerobic actinomycetes in sterile distilled water. Appl. Microbiol. 28:218-222.
Ryan, M. J., P. D. Bridge, D. Smith, and P. Jeffries. 2002. Phenotypic degeneration occurs during sector formation in Metarhizium anisopliae. J. Appl. Microbiol. 93:163-168.
Ryan, M. J., P. Jeffries, P. D. Bridge, and D. Smith. 2001. Developing cryopreservation protocols to secure fungal gene function. Cryo Lett. 22:115-124.
East Kent Microbiology Service, The William Harvey Hospital, Kennington Road, Ashford, Kent TN24 0LZ
1 Department of Biosciences, University of Kent, Canterbury, Kent CT2 6NJ, United Kingdom(M. Baker and P. Jeffries)