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Identification of mollicutes from insects
D12 IDENTIFICATION OF MOLLICUTES FROM INSECTS Robert F. Whitcomb and Kevin j . Hackett
Introduction After the initial discovery that spiroplasmas were associated principally with arthropods, major efforts were made to isolate mollicutes from diverse insect taxa and from ecosystems that the insects inhabited (Hackett and Clark, 1989). These studies resulted in the discovery of a large number of Spiroplasma species and several additional genera of mollicutes. One of these genera (Acholeplasma) appears to inhabit various niches in nature, including vertebrate animals, insects, and plant surfaces. In rare cases, members of the genus Mycoplasma have apparently been isolated from plants and insects, as have some L-phase variants of bacteria. The latter two types of wall-less prokaryotes are probably not associated with arthropods, but have been encountered as artifacts of the isolation procedure. Other taxa have proved to be predictably associated with arthropods, especially insects. These taxa have formed the basis for a new order of arthropod-associated mollicutes, the Entomoplasmatales (Tully et al., 1993). Techniques for isolation of mollicutes from insects (Markham et al., 1983; Hackett and Clark, 1989) and for preparation of media suitable for isolations from these habitats are reviewed in Volume I of this series (see Chapters A2, A3, and A4) and in chapter DIO of this volume. This chapter describes the identification procedures for wall-less prokaryotes isolated from arthropods.
313 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Robert F. Whitcomb and Kevin J. Hackett
Steps to Be Taken before Identification Begins If only a few isolates are obtained in a given study, it is likely that each will be important enough to merit detailed study. In this case, each isolate should be cloned (see Chapter E2 in Vol. I). Motile, helical organisms that are absolutely resistant to penicillin can be tentatively presumed to be spiroplasmas. If they turn out to represent a new group or subgroup, the lack of cell wall must be confirmed by electron microscopy (see Chapter E2 in Vol. I) in the course of description of the new group (Whitcomb et al., 1987). If the organisms are nonhelical, some consideration should be given to the possibility that they could be bacterial L-phase variants (see chapter E2, Vol. I). If this is deemed unlikely, the organisms should be passed to a bank of diagnostic media, including serum-containing and serum-free media with or without a polysorbitan monooleate (Tween 80) supplement (see Chapter E7, Vol. I). From these preliminary tests, the genus can be tentatively assigned. In some studies, the isolation rate may be so high that it is not feasible to clone every isolate. If this is true, the possibility that diagnostic tests have been performed on a mixed culture must be taken into account. In the section on identification by the spiroplasma deformation test (see the following section), the detection of mixtures is discussed.
Identification of Genera of Wall-less Mollicutes from Insects Identification of mollicutes from insects has become increasingly complex; new genera of organisms have been discovered in arthropods and species and groups from the diverse genera have proliferated. The following key gives a guide to the principles involved in distinguishing the genera of mollicutes associated with arthropods. Key to Genera of Wall-less Prokaryotes from Arthropods 1. 2.
3. 4.
Helical Spiroplasma Nonhelical 2 Passes 450-(xm but not 220-|jLm filter; reverts to walled bacterium when passed in conventional antibiotic-free bacteriological media Bacterial L-phase variant Filterable through 220-|xm filters; penicillin-resistant; verified by electron microscopy to be wall-less 3 Grows in serum-free mycoplasma broth Acholeplasma Does not grow in serum-free broth 4 Grows in serum-free broth supplemented with 0.04% Tween 80 Mesoplasma Does not grow in serum-free broth with Tween 80 5
D12 Identifying Mollicutes from Insects 5.
6.
315
Reverts to helical morphology at some stage in growth cycle when tested with various media Spiroplasma Does not revert 6 Grows optimally at 37°C, possible contaminant from serum in medium or from the ecosystem Mycoplasma Grows optimally at a temperature less than 37°C, often 30° or 32°C; may grow at 20°C or less Entomoplasma
Identification to Species or Group Level Although the species is the central unit in mollicute taxonomy, it is an arbitrarily defined unit. Several features characterize the species. The first is that the clusters of strains that constitute mollicute species differ from other clusters by at least 30% of their genome (i.e., there is less than 70% DNA-DNA homology as measured by hybridization studies). Description of new species must conform to minimal standards (see Chapter E2 in Vol. I). The species (or, in the case of spiroplasmas, the group) is determined, as a practical matter, by serology. When the genus of a strain has been determined, the culture should be tested against specific antisera directed to the known species of that genus. Nonhelical organisms should be tested by the growth inhibition test (Tully et aL, 1994). Helical organisms should be tested by the spiroplasma deformation test, as detailed later. Antisera are available, at present, to 12 Mesoplasma, 5 Entomoplasma, and 13 Acholeplasma species. In the unlikely event that an isolate should turn out to be a true Mycoplasma species, it must be tested against sera to about 100 species. If the organism is helical, sera are available for 25 groups (17 of which have been given binomial names), 11 unique subgroups, and 8 putative groups that are well characterized (Tully et aL, 1987; Whitcomb et aL, 1992b; See also Appendix, this volume). In tests with particular Spiroplasma or Mycoplasma species, it is sometimes important to use antiserum to one or two other strains within the species complex. This ensures that the test spans the antigenic variability of the species; failure to do so could result in missing the identity of the unknown organism. Species and strain designations of mollicutes are given in the Appendix to this volume.
Identification of Infraspecific Units Various hierarchies of variation have emerged from the detailed study of mollicute strains (Whitcomb, 1994). In spiroplasmas, the group is a cluster of strains that is serologically unrelated to all other groups. In cases where DNA-
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DNA homology data are available, groups have turned out to share little or no homology; thus the spiroplasma serogroups can be thought of, tentatively, as homology groups. The term subgroup has been used to denote clusters of strains that share a considerable amount of DNA-DNA homology, but that are nonetheless substantially different (Whitcomb et al., 1987). The subspecies designation in mollicutes has been reserved for special cases in which it has been desirable to construct a formal taxonomy for economically important clusters of strains that share more than 70% DNA-DNA homology, but less than about 80%. In many spiroplasma assemblages (e.g., those from horseflies and mosquitoes, and the various representatives of group IV spiroplasmas from a number of sources), a wide range of variability has been encountered. Variability in these groups, although less than that between subgroups, is still substantial and appears to reflect different habitats and ecologies from which the variants have been derived. Such variation can be described at the serovar level. Spiroplasma serovars described by deformation tests can be described in terms of their reactivity with several antisera (see the next section). In many cases, there will be a lack of background information on the meaning of partial serological reactions. If such is the case, identifications will of necessity be only partial; many isolates may be identified as being serologically related to the representative strain of a known group, but not identical to it. A discussion of the issues involved in infraspecific variation is beyond the scope of this chapter, but is presented in several reports (Whitcomb et al, 1987; Whitcomb, 1994).
Spiroplasma Deformation Test Adaptation of the spiroplasma deformation test to routine diagnosis has resulted in significant modifications to previously reported protocols (Williamson and Whitcomb, 1983). We present the protocol for the test as currently performed. Materials Culture of an unknown spiroplasma in logarithmic growth phase in MID medium (Whitcomb, 1983); some tick-associated spiroplasmas may require SP-4 medium (see Chapter A2 in Vol. I) Dilutions of antiserum specific to each group (25 sera), subgroup (11 sera), and putative group (8 sera), in twofold dilutions (1:10 to 1:20,480) in MID medium Polyvalent antisera designed in accordance with past cultural experience 96-well microtiter plates
D12 Identifying Mollicutes from insects
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Automatic pipetter Dark-field microscope
Procedure ANTIGEN-ANTIBODY REACTION
1. Pipette 25 jxl of each of the 12 antiserum dilutions into a single row of wells in the microtiter plates. 2. Adjust antiserum dilutions to about 10^ helical cells/ml (Rodwell and Whitcomb, 1983). 3. Add 25 fxl antigen to each antiserum dilution. 4. After 30 minutes, examine antigen-antibody mixtures under dark-field microscope. CHOICE OF TEST SERA
In many cases, a background of previous data produces an expectation of isolation for certain spiroplasma groups and/or species. When this is true, it is possible to screen isolates against a reduced bank of antisera and to further reduce the work volume by use of polyvalent sera constructed by mixing univalent, independently produced antisera. For example, six spiroplasma groups are regularly isolated from horseflies in the southeastern United States (French et al., 1990; Whitcomb et al., 1992a). We screen new isolates against a bank of only five polyvalent sera. One of these contains antibodies at a final dilution of 1:10 to the B31T (S. apis), W13, and PPSl strains of group IV; although this group appears in tabanids in northern latitudes and in Europe (Le Goff et al., 1991), it is very rare in tabanids in the southeastern United States. The second polyvalent antiserum contains antisera to the three representative strains of the subgroups of spiroplasma group VIII: strains EA-1 (VIII-1), DF-1 (VIII-2), and TAAS-1 (VIII-3) (Gasparich et al., 1993). The third polyvalent serum contains sera directed against the group XIV EC-1 strain and the ungrouped HYOS-1 strain. The fourth polyvalent serum contains antibodies directed against the group XXIII strain TG-1 and the ungrouped TAUS-1 strain. Most strains from the southeast react with one of the last three polyvalent sera. Occasionally, new isolates react with polyvalent serum number 5, which contains a mixture of antibodies, each a dilution of 1:10, against the group XVIII TN-1 strain and ungrouped strains TALS-2 and TABS-2; each of the latter two strains is a candidate for status as a new spiroplasma group. This typing scheme is adequate for tabanid spiroplasma assemblages from the southeastern United States. In a different geographical region, the experience may be quite different. Strains
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^!/,f'A!
Fig. 1. Diagrammatic representation of various microscopic observations obtained in spiroplasma deformation tests carried out under dark-field microscopy. (A) In the absence of a specific antibody, the culture consists predominantly of helical, motile cells; (B) end point of a deformation test, at which approximately one-half of the cells are deformed and exhibit blebs; (C) high levels of antibody induce immobilization and multiple blebbing, vi'ith agglutination of affected cells; and (D) evidence of mixed culture where the field contains some motile and helical unaffected organisms, but also aggregated and multiply blebbed cells.
isolated from horseflies in Europe exhibit a very different profile (Le Goff et al., 1991; C. Chastel, personal communication). READING THE TEST
1. If the culture contains largely typical, helical motile spiroplasma cells (Fig. lA), the test can be read as previously described (Williamson and Whitcomb, 1983). Briefly, the dilution at which about one-half of the helical cells have been deformed (Fig. IB) is recorded, and the titer of the reaction is expressed as the reciprocal of the end-point dilution.
D12 Identifying Mollicutes from Insects
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il^
Fig. 1. {Continued)
2. In many primary cultures, all or many spiroplasma cells are not perfect helices. It is common for many cells to be filamentous, to have a certain amount of nonspecific blebbing, or to have many very short or almost completely nonhelical forms. In all of these cases, it is possible to recognize serological deformation, providing a control culture that has not reacted with a specific antibody is available. In most cases, the well with the lowest antibody-antigen ratio (i.e., well No. 12, total dilution 1:41,960) will serve as this control. In a few cases, in which serological reactions may occur at very dilute antiserum concentrations, it may be necessary to use a control consisting of cultured organisms to which an equal volume of medium has been added. Ideally, in this control, there will be a large population of unaffected cells (Fig. 1 A). If more than one set of antibodies is utilized, the negative reactions from antisera that do not react with the unknown spiroplasma serve as auxilliary controls; these reactions control, to some extent, for the possible effect of a nonhomologous antibody and other components that may be present in heterologous antisera.
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3. In reading the results, it is important to be thoroughly familiar with the various types of spiroplasma-antibody reactions. At very high levels of antibody attack, some spiroplasmas (e.g., S. clarkii) are immobilized without blebbing; it would be possible to mistake this response for a negative reaction. At high levels of antibody attack, the aspect of the affected cells is one of multiply blebbed cells, with varying degrees of agglutination between affected cells (Fig. IC). As a result, the total number of bodies in the reaction mixture is greatly reduced, and no free helical filament, attached or unattached to any bleb or bleb mass, is found in the culture. As the ratio of antibody with respect to antigen decreases, a point is reached at which some cells have portions of helical filaments that are unaffected by antibody. This is true, even though 100% of the cells may show some effect of antibody. At somewhat lower levels of antibody, some cells escape attack altogether; eventually, the fraction of affected cells decreases to about half—this is the end point. Affected cells, although constituting less than 50% of the population, are usually evident at antibody attack rates one-fourth or, rarely, one-eighth those at the end point. 4. Although mixed infections are always of some concern in series of primary isolations, certain hosts carry frequent infections of different spiroplasmas. Such cases offer serious challenges in serotyping of primary isolates. In other mollicute systems, in which primary isolations are carried out on solid media, mixtures can be readily detected by use of fluorescent antibody techniques (see Chapter B9, this volume). Because spiroplasmas produce colonies erratically if at all, few if any spiroplasmas are isolated on solid media. The problem is, then, to recognize a mixture during primary isolation in liquid media. This has been accomplished in our laboratory by noting the morphology of affected spiroplasmas. The reaction series of an unmixed isolate, or of a culture that has a great predominance of a single spiroplasma serovar, is usually seen as a progression, from aggregated multiply blebbed cells, to populations of cells, some of which have single blebs. The identity of such cultures can be tentatively established by normal deformation tests. In contrast, the presence in a single antigen-antibody reaction of aggregated multiply blebbed cells and free, unaffected cells (Fig. ID) is a clear warning of a mixed culture. If such a culture is passed 5-10 times, the serological profile usually shifts, as one of the components of the original mixture eventually outgrows the other. Thus, a shift in apparent serological identity during cultural history is a second warning of a mixed culture. Resolution of such a mixture to identify all spiroplasmas may be very difficult, especially if one or both of the spiroplasmas react only partially with the test sera. INTERPRETATION OF RESULTS
Interpretation depends on the degree of knowledge about the mollicutes associated with the given host. Identification to the group level is usually not prob-
D12 Identifying Mollicutes from Insects
321
lematic, if the unknown culture has a significant reaction to existing group antisera. Further testing, once preUminary screening has been carried out, consists of setting up complete dilution series with each of the components of the polyvalent sera. The final titer is determined by these tests. In many cases with tabanid spiroplasmas, some intergroup cross-reactivity, especially one-way crosses, has been observed. This circumstance, while unfortunate in one sense, can be exploited in defining spiroplasma serovars. For example, we always record titers of group VIII strains against all three representative group VIII sera. The set of three titers defines a level of reactivity that defines common serovars that occur in Georgia (French et al, 1990; Whitcomb et al., 1992a). Similarly, strains that react to polyvalent sera numbers 3 or 4 are tested against all four of the component sera of these polyvalent reagents. The matrix of titers obtained, as with group VIII, establishes four titers for each isolate and is a sensitive means of establishing serovar identity.
Discussion Mollicutes abound in arthropods, especially in insects. The meaning of infraspecific mollicute clusters from these vast reservoirs is not yet clear, but has formed a rich basis for speculation about the role that habitat, natural selection, and genetic drift play in the determination of mollicute serovars (Whitcomb, 1994). Identification of a large bank of strains related at various hierarchical levels will contribute to an understanding of the natural ecology of mollicutes in their insect hosts.
References French, F. E., Whitcomb, R. F., Tully, J. G., Hackett, K. J., Clark, E. A., Henegar, R. B., and Rose, D. L. (1990). Tabanid spiroplasmas of the southeastern USA: New groups, and correlation with host life history strategy. Zentralbl. BakterioL, Suppl. 20, 441-444. Gasparich, G., Whitcomb, R. F., French, F. E., Clark, E. A., and Tully, J. G. (1993). Serologic and genomic relatedness of group VIII and group XVII spiroplasmas and subdivision of spiroplasma group VIII into subgroups. Int. J. Syst. Bacteriol. 43, 338-341. Hackett, K. J., and Clark, T. B. (1989). Ecology of spiroplasmas. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 113-200. Academic Press, San Diego, CA. Le Goff, F., Humphery-Smith, I., Leclercq, M., and Chastel, C. (1991). Spiroplasmas from European Tabanidae. Med. Vet. Entomol. 5, 143-144. Markham, P., Clark, T. B., and Whitcomb, R. F. (1983). Culture techniques for spiroplasmas from arthropods. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 217-223. Academic Press, New York. Rodwell, A., and Whitcomb, R. F. (1983). Methods for direct and indirect measurement of myco-
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plasma growth. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 185-196. Academic Press, New York. Tully, J. G., Rose, D. L., Clark, E., Carle, P., Bove, J. M., Whitcomb, R. F., Colflesh, D. E., Henegar, R. B., and Williamson, D. L. (1987). Revised group classification of the genus Spiroplasma (class Mollicutes) with proposed new groups XII to XXII. Int. J. Syst. Bacteriol. 31, 357-364. Tully, J. G., Bove, J. M., Laigret, P., and Whitcomb, R. F. (1993). Revised taxonomy of the class Mollicutes: Proposed elevation of a monophyletic cluster of arthropod-associated mollicutes to ordinal rank (Entomoplasmatales ord. nov.), with provision for familial rank to separate species with nonhelical morphology (Entomoplasmataceae fam. nov.) from helical species {Spiroplasmataceae), and emended description of the order Mycoplasmatales, family Mycoplasmataceae. Int. J. Syst. Bacteriol. 43, 378-385. Tully, J. G., Whitcomb, R. F., Hackett, K. J., Rose, D. L., Henegar, R. B., Bove, J. M., Carle, P., Williamson, D. L., and Clark, T. B. (1994). Taxonomic descriptions of eight new non-sterolrequiring mollicutes assigned to the genus Mesoplasma. Int. J. Syst. Bacteriol. 44, 685-693. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.) Vol. 1, pp. 147-158. Academic Press, New York. Whitcomb, R. F. (1994). The species concept in eukaryotes and prokaryotes: Search for a synthesis. lOM Lett. 3, 1-7. Whitcomb, R. F., Bove, J. M., Chen, T. A., Tully, J. G., and Williamson, D. L. (1987). Proposed criteria for an interim serogroup classification for members of the genus Spiroplasma (Class Mollicutes). Int. J. Syst. Bacteriol. 37, 82-84. Whitcomb, R. F., French, F. E., Tully, J. G., Gasparich, G. E., Bove, J. M., Carle, P., Clark, E. A., and Henegar, R. (1992a). Tabanid spiroplasma serovars. lOM Lett. 2, 115. Whitcomb, R. F., Tully, J. G., Williamson, D. L., Bove, J. M., French, F. E., Konai, M., Gasparich, G., Abalain-Colloc, M. L., Saillard, C , Chastel, C , Carle, P., Rose, D. L., Henegar, R., Clark, E. A., and Hackett, K. J. (1992b). Revised classification of spiroplasmas. lOMLett. 2, 134. Williamson, D. L., and Whitcomb, R. F. (1983). Special serological tests for spiroplasma identification. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 249-259. Academic Press, New York.
Introduction After the initial discovery that spiroplasmas were associated principally with arthropods, major efforts were made to isolate mollicutes from diverse insect taxa and from ecosystems that the insects inhabited (Hackett and Clark, 1989). These studies resulted in the discovery of a large number of Spiroplasma species and several additional genera of mollicutes. One of these genera (Acholeplasma) appears to inhabit various niches in nature, including vertebrate animals, insects, and plant surfaces. In rare cases, members of the genus Mycoplasma have apparently been isolated from plants and insects, as have some L-phase variants of bacteria. The latter two types of wall-less prokaryotes are probably not associated with arthropods, but have been encountered as artifacts of the isolation procedure. Other taxa have proved to be predictably associated with arthropods, especially insects. These taxa have formed the basis for a new order of arthropod-associated mollicutes, the Entomoplasmatales (Tully et al., 1993). Techniques for isolation of mollicutes from insects (Markham et al., 1983; Hackett and Clark, 1989) and for preparation of media suitable for isolations from these habitats are reviewed in Volume I of this series (see Chapters A2, A3, and A4) and in chapter DIO of this volume. This chapter describes the identification procedures for wall-less prokaryotes isolated from arthropods.
313 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Steps to Be Taken before Identification Begins If only a few isolates are obtained in a given study, it is likely that each will be important enough to merit detailed study. In this case, each isolate should be cloned (see Chapter E2 in Vol. I). Motile, helical organisms that are absolutely resistant to penicillin can be tentatively presumed to be spiroplasmas. If they turn out to represent a new group or subgroup, the lack of cell wall must be confirmed by electron microscopy (see Chapter E2 in Vol. I) in the course of description of the new group (Whitcomb et al., 1987). If the organisms are nonhelical, some consideration should be given to the possibility that they could be bacterial L-phase variants (see chapter E2, Vol. I). If this is deemed unlikely, the organisms should be passed to a bank of diagnostic media, including serum-containing and serum-free media with or without a polysorbitan monooleate (Tween 80) supplement (see Chapter E7, Vol. I). From these preliminary tests, the genus can be tentatively assigned. In some studies, the isolation rate may be so high that it is not feasible to clone every isolate. If this is true, the possibility that diagnostic tests have been performed on a mixed culture must be taken into account. In the section on identification by the spiroplasma deformation test (see the following section), the detection of mixtures is discussed.
Identification of Genera of Wall-less Mollicutes from Insects Identification of mollicutes from insects has become increasingly complex; new genera of organisms have been discovered in arthropods and species and groups from the diverse genera have proliferated. The following key gives a guide to the principles involved in distinguishing the genera of mollicutes associated with arthropods. Key to Genera of Wall-less Prokaryotes from Arthropods 1. 2.
3. 4.
Helical Spiroplasma Nonhelical 2 Passes 450-(xm but not 220-|jLm filter; reverts to walled bacterium when passed in conventional antibiotic-free bacteriological media Bacterial L-phase variant Filterable through 220-|xm filters; penicillin-resistant; verified by electron microscopy to be wall-less 3 Grows in serum-free mycoplasma broth Acholeplasma Does not grow in serum-free broth 4 Grows in serum-free broth supplemented with 0.04% Tween 80 Mesoplasma Does not grow in serum-free broth with Tween 80 5
D12 Identifying Mollicutes from Insects 5.
6.
315
Reverts to helical morphology at some stage in growth cycle when tested with various media Spiroplasma Does not revert 6 Grows optimally at 37°C, possible contaminant from serum in medium or from the ecosystem Mycoplasma Grows optimally at a temperature less than 37°C, often 30° or 32°C; may grow at 20°C or less Entomoplasma
Identification to Species or Group Level Although the species is the central unit in mollicute taxonomy, it is an arbitrarily defined unit. Several features characterize the species. The first is that the clusters of strains that constitute mollicute species differ from other clusters by at least 30% of their genome (i.e., there is less than 70% DNA-DNA homology as measured by hybridization studies). Description of new species must conform to minimal standards (see Chapter E2 in Vol. I). The species (or, in the case of spiroplasmas, the group) is determined, as a practical matter, by serology. When the genus of a strain has been determined, the culture should be tested against specific antisera directed to the known species of that genus. Nonhelical organisms should be tested by the growth inhibition test (Tully et aL, 1994). Helical organisms should be tested by the spiroplasma deformation test, as detailed later. Antisera are available, at present, to 12 Mesoplasma, 5 Entomoplasma, and 13 Acholeplasma species. In the unlikely event that an isolate should turn out to be a true Mycoplasma species, it must be tested against sera to about 100 species. If the organism is helical, sera are available for 25 groups (17 of which have been given binomial names), 11 unique subgroups, and 8 putative groups that are well characterized (Tully et aL, 1987; Whitcomb et aL, 1992b; See also Appendix, this volume). In tests with particular Spiroplasma or Mycoplasma species, it is sometimes important to use antiserum to one or two other strains within the species complex. This ensures that the test spans the antigenic variability of the species; failure to do so could result in missing the identity of the unknown organism. Species and strain designations of mollicutes are given in the Appendix to this volume.
Identification of Infraspecific Units Various hierarchies of variation have emerged from the detailed study of mollicute strains (Whitcomb, 1994). In spiroplasmas, the group is a cluster of strains that is serologically unrelated to all other groups. In cases where DNA-
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DNA homology data are available, groups have turned out to share little or no homology; thus the spiroplasma serogroups can be thought of, tentatively, as homology groups. The term subgroup has been used to denote clusters of strains that share a considerable amount of DNA-DNA homology, but that are nonetheless substantially different (Whitcomb et al., 1987). The subspecies designation in mollicutes has been reserved for special cases in which it has been desirable to construct a formal taxonomy for economically important clusters of strains that share more than 70% DNA-DNA homology, but less than about 80%. In many spiroplasma assemblages (e.g., those from horseflies and mosquitoes, and the various representatives of group IV spiroplasmas from a number of sources), a wide range of variability has been encountered. Variability in these groups, although less than that between subgroups, is still substantial and appears to reflect different habitats and ecologies from which the variants have been derived. Such variation can be described at the serovar level. Spiroplasma serovars described by deformation tests can be described in terms of their reactivity with several antisera (see the next section). In many cases, there will be a lack of background information on the meaning of partial serological reactions. If such is the case, identifications will of necessity be only partial; many isolates may be identified as being serologically related to the representative strain of a known group, but not identical to it. A discussion of the issues involved in infraspecific variation is beyond the scope of this chapter, but is presented in several reports (Whitcomb et al, 1987; Whitcomb, 1994).
Spiroplasma Deformation Test Adaptation of the spiroplasma deformation test to routine diagnosis has resulted in significant modifications to previously reported protocols (Williamson and Whitcomb, 1983). We present the protocol for the test as currently performed. Materials Culture of an unknown spiroplasma in logarithmic growth phase in MID medium (Whitcomb, 1983); some tick-associated spiroplasmas may require SP-4 medium (see Chapter A2 in Vol. I) Dilutions of antiserum specific to each group (25 sera), subgroup (11 sera), and putative group (8 sera), in twofold dilutions (1:10 to 1:20,480) in MID medium Polyvalent antisera designed in accordance with past cultural experience 96-well microtiter plates
D12 Identifying Mollicutes from insects
317
Automatic pipetter Dark-field microscope
Procedure ANTIGEN-ANTIBODY REACTION
1. Pipette 25 jxl of each of the 12 antiserum dilutions into a single row of wells in the microtiter plates. 2. Adjust antiserum dilutions to about 10^ helical cells/ml (Rodwell and Whitcomb, 1983). 3. Add 25 fxl antigen to each antiserum dilution. 4. After 30 minutes, examine antigen-antibody mixtures under dark-field microscope. CHOICE OF TEST SERA
In many cases, a background of previous data produces an expectation of isolation for certain spiroplasma groups and/or species. When this is true, it is possible to screen isolates against a reduced bank of antisera and to further reduce the work volume by use of polyvalent sera constructed by mixing univalent, independently produced antisera. For example, six spiroplasma groups are regularly isolated from horseflies in the southeastern United States (French et al., 1990; Whitcomb et al., 1992a). We screen new isolates against a bank of only five polyvalent sera. One of these contains antibodies at a final dilution of 1:10 to the B31T (S. apis), W13, and PPSl strains of group IV; although this group appears in tabanids in northern latitudes and in Europe (Le Goff et al., 1991), it is very rare in tabanids in the southeastern United States. The second polyvalent antiserum contains antisera to the three representative strains of the subgroups of spiroplasma group VIII: strains EA-1 (VIII-1), DF-1 (VIII-2), and TAAS-1 (VIII-3) (Gasparich et al., 1993). The third polyvalent serum contains sera directed against the group XIV EC-1 strain and the ungrouped HYOS-1 strain. The fourth polyvalent serum contains antibodies directed against the group XXIII strain TG-1 and the ungrouped TAUS-1 strain. Most strains from the southeast react with one of the last three polyvalent sera. Occasionally, new isolates react with polyvalent serum number 5, which contains a mixture of antibodies, each a dilution of 1:10, against the group XVIII TN-1 strain and ungrouped strains TALS-2 and TABS-2; each of the latter two strains is a candidate for status as a new spiroplasma group. This typing scheme is adequate for tabanid spiroplasma assemblages from the southeastern United States. In a different geographical region, the experience may be quite different. Strains
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^!/,f'A!
Fig. 1. Diagrammatic representation of various microscopic observations obtained in spiroplasma deformation tests carried out under dark-field microscopy. (A) In the absence of a specific antibody, the culture consists predominantly of helical, motile cells; (B) end point of a deformation test, at which approximately one-half of the cells are deformed and exhibit blebs; (C) high levels of antibody induce immobilization and multiple blebbing, vi'ith agglutination of affected cells; and (D) evidence of mixed culture where the field contains some motile and helical unaffected organisms, but also aggregated and multiply blebbed cells.
isolated from horseflies in Europe exhibit a very different profile (Le Goff et al., 1991; C. Chastel, personal communication). READING THE TEST
1. If the culture contains largely typical, helical motile spiroplasma cells (Fig. lA), the test can be read as previously described (Williamson and Whitcomb, 1983). Briefly, the dilution at which about one-half of the helical cells have been deformed (Fig. IB) is recorded, and the titer of the reaction is expressed as the reciprocal of the end-point dilution.
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il^
Fig. 1. {Continued)
2. In many primary cultures, all or many spiroplasma cells are not perfect helices. It is common for many cells to be filamentous, to have a certain amount of nonspecific blebbing, or to have many very short or almost completely nonhelical forms. In all of these cases, it is possible to recognize serological deformation, providing a control culture that has not reacted with a specific antibody is available. In most cases, the well with the lowest antibody-antigen ratio (i.e., well No. 12, total dilution 1:41,960) will serve as this control. In a few cases, in which serological reactions may occur at very dilute antiserum concentrations, it may be necessary to use a control consisting of cultured organisms to which an equal volume of medium has been added. Ideally, in this control, there will be a large population of unaffected cells (Fig. 1 A). If more than one set of antibodies is utilized, the negative reactions from antisera that do not react with the unknown spiroplasma serve as auxilliary controls; these reactions control, to some extent, for the possible effect of a nonhomologous antibody and other components that may be present in heterologous antisera.
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3. In reading the results, it is important to be thoroughly familiar with the various types of spiroplasma-antibody reactions. At very high levels of antibody attack, some spiroplasmas (e.g., S. clarkii) are immobilized without blebbing; it would be possible to mistake this response for a negative reaction. At high levels of antibody attack, the aspect of the affected cells is one of multiply blebbed cells, with varying degrees of agglutination between affected cells (Fig. IC). As a result, the total number of bodies in the reaction mixture is greatly reduced, and no free helical filament, attached or unattached to any bleb or bleb mass, is found in the culture. As the ratio of antibody with respect to antigen decreases, a point is reached at which some cells have portions of helical filaments that are unaffected by antibody. This is true, even though 100% of the cells may show some effect of antibody. At somewhat lower levels of antibody, some cells escape attack altogether; eventually, the fraction of affected cells decreases to about half—this is the end point. Affected cells, although constituting less than 50% of the population, are usually evident at antibody attack rates one-fourth or, rarely, one-eighth those at the end point. 4. Although mixed infections are always of some concern in series of primary isolations, certain hosts carry frequent infections of different spiroplasmas. Such cases offer serious challenges in serotyping of primary isolates. In other mollicute systems, in which primary isolations are carried out on solid media, mixtures can be readily detected by use of fluorescent antibody techniques (see Chapter B9, this volume). Because spiroplasmas produce colonies erratically if at all, few if any spiroplasmas are isolated on solid media. The problem is, then, to recognize a mixture during primary isolation in liquid media. This has been accomplished in our laboratory by noting the morphology of affected spiroplasmas. The reaction series of an unmixed isolate, or of a culture that has a great predominance of a single spiroplasma serovar, is usually seen as a progression, from aggregated multiply blebbed cells, to populations of cells, some of which have single blebs. The identity of such cultures can be tentatively established by normal deformation tests. In contrast, the presence in a single antigen-antibody reaction of aggregated multiply blebbed cells and free, unaffected cells (Fig. ID) is a clear warning of a mixed culture. If such a culture is passed 5-10 times, the serological profile usually shifts, as one of the components of the original mixture eventually outgrows the other. Thus, a shift in apparent serological identity during cultural history is a second warning of a mixed culture. Resolution of such a mixture to identify all spiroplasmas may be very difficult, especially if one or both of the spiroplasmas react only partially with the test sera. INTERPRETATION OF RESULTS
Interpretation depends on the degree of knowledge about the mollicutes associated with the given host. Identification to the group level is usually not prob-
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lematic, if the unknown culture has a significant reaction to existing group antisera. Further testing, once preUminary screening has been carried out, consists of setting up complete dilution series with each of the components of the polyvalent sera. The final titer is determined by these tests. In many cases with tabanid spiroplasmas, some intergroup cross-reactivity, especially one-way crosses, has been observed. This circumstance, while unfortunate in one sense, can be exploited in defining spiroplasma serovars. For example, we always record titers of group VIII strains against all three representative group VIII sera. The set of three titers defines a level of reactivity that defines common serovars that occur in Georgia (French et al, 1990; Whitcomb et al., 1992a). Similarly, strains that react to polyvalent sera numbers 3 or 4 are tested against all four of the component sera of these polyvalent reagents. The matrix of titers obtained, as with group VIII, establishes four titers for each isolate and is a sensitive means of establishing serovar identity.
Discussion Mollicutes abound in arthropods, especially in insects. The meaning of infraspecific mollicute clusters from these vast reservoirs is not yet clear, but has formed a rich basis for speculation about the role that habitat, natural selection, and genetic drift play in the determination of mollicute serovars (Whitcomb, 1994). Identification of a large bank of strains related at various hierarchical levels will contribute to an understanding of the natural ecology of mollicutes in their insect hosts.
References French, F. E., Whitcomb, R. F., Tully, J. G., Hackett, K. J., Clark, E. A., Henegar, R. B., and Rose, D. L. (1990). Tabanid spiroplasmas of the southeastern USA: New groups, and correlation with host life history strategy. Zentralbl. BakterioL, Suppl. 20, 441-444. Gasparich, G., Whitcomb, R. F., French, F. E., Clark, E. A., and Tully, J. G. (1993). Serologic and genomic relatedness of group VIII and group XVII spiroplasmas and subdivision of spiroplasma group VIII into subgroups. Int. J. Syst. Bacteriol. 43, 338-341. Hackett, K. J., and Clark, T. B. (1989). Ecology of spiroplasmas. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 113-200. Academic Press, San Diego, CA. Le Goff, F., Humphery-Smith, I., Leclercq, M., and Chastel, C. (1991). Spiroplasmas from European Tabanidae. Med. Vet. Entomol. 5, 143-144. Markham, P., Clark, T. B., and Whitcomb, R. F. (1983). Culture techniques for spiroplasmas from arthropods. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 217-223. Academic Press, New York. Rodwell, A., and Whitcomb, R. F. (1983). Methods for direct and indirect measurement of myco-
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plasma growth. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 185-196. Academic Press, New York. Tully, J. G., Rose, D. L., Clark, E., Carle, P., Bove, J. M., Whitcomb, R. F., Colflesh, D. E., Henegar, R. B., and Williamson, D. L. (1987). Revised group classification of the genus Spiroplasma (class Mollicutes) with proposed new groups XII to XXII. Int. J. Syst. Bacteriol. 31, 357-364. Tully, J. G., Bove, J. M., Laigret, P., and Whitcomb, R. F. (1993). Revised taxonomy of the class Mollicutes: Proposed elevation of a monophyletic cluster of arthropod-associated mollicutes to ordinal rank (Entomoplasmatales ord. nov.), with provision for familial rank to separate species with nonhelical morphology (Entomoplasmataceae fam. nov.) from helical species {Spiroplasmataceae), and emended description of the order Mycoplasmatales, family Mycoplasmataceae. Int. J. Syst. Bacteriol. 43, 378-385. Tully, J. G., Whitcomb, R. F., Hackett, K. J., Rose, D. L., Henegar, R. B., Bove, J. M., Carle, P., Williamson, D. L., and Clark, T. B. (1994). Taxonomic descriptions of eight new non-sterolrequiring mollicutes assigned to the genus Mesoplasma. Int. J. Syst. Bacteriol. 44, 685-693. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.) Vol. 1, pp. 147-158. Academic Press, New York. Whitcomb, R. F. (1994). The species concept in eukaryotes and prokaryotes: Search for a synthesis. lOM Lett. 3, 1-7. Whitcomb, R. F., Bove, J. M., Chen, T. A., Tully, J. G., and Williamson, D. L. (1987). Proposed criteria for an interim serogroup classification for members of the genus Spiroplasma (Class Mollicutes). Int. J. Syst. Bacteriol. 37, 82-84. Whitcomb, R. F., French, F. E., Tully, J. G., Gasparich, G. E., Bove, J. M., Carle, P., Clark, E. A., and Henegar, R. (1992a). Tabanid spiroplasma serovars. lOM Lett. 2, 115. Whitcomb, R. F., Tully, J. G., Williamson, D. L., Bove, J. M., French, F. E., Konai, M., Gasparich, G., Abalain-Colloc, M. L., Saillard, C , Chastel, C , Carle, P., Rose, D. L., Henegar, R., Clark, E. A., and Hackett, K. J. (1992b). Revised classification of spiroplasmas. lOMLett. 2, 134. Williamson, D. L., and Whitcomb, R. F. (1983). Special serological tests for spiroplasma identification. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 249-259. Academic Press, New York.