Antibiotics in Laboratory Medicine, 6 Ed.

Chapter 7. Antimicrobial Susceptibility Testing for Some Atypical Microorganisms (Chlamydia, Mycoplasma, Rickettsia, Ehrlichia, Coxiella, and Spirochetes)

Jean-Marc Rolain

This chapter discusses susceptibility testing for fastidious organisms, including mycoplasmas (1), Borrelia burgdorferi (2), Leptospira, and those that cannot be cultured without the use of animals or tissue culture (Chlamydia, Rickettsia, Ehrlichia, and Coxiella burnetii). The special testing requirements using cells make it very difficult for all but research laboratories to perform susceptibility testing of these organisms. Thus, the susceptibility testing that has been done has been somewhat limited in terms of both the number of isolates tested and the number of different antiinfective agents evaluated. Furthermore, frequently, only well-characterized laboratory strains have been tested rather than recent clinical isolates.

Nonetheless, continual progress is being made in the use of tissue culture for some of these highly fastidious agents, such as Treponema pallidum (3), and it should soon be possible to carry out more extensive studies. Expanding the range of studies will certainly be important, because resistance will continue to remain unconfirmed if testing is not carried out (4). This will be true even if resistance is suspected from observations of patients, as was true in the case of a T. pallidum infection (135).

The complexity of the methods needed to propagate these organisms argues for an attempt to standardize the methods used for susceptibility testing. In the case of chlamydiae, standardization should be possible, but despite the many different techniques used for the testing of chlamydiae, the results obtained have been remarkably consistent (6). Nonetheless, greater uniformity of testing techniques would make it easier to compare results obtained in different laboratories. European guidelines for susceptibility testing of intracellular and cell-associated pathogens have been recently published and should be used by laboratories to allow comparison of the results obtained (7).

Since the previous publication of this chapter, new information has been published on the susceptibility of chlamydiae, mycoplasmas, and rickettsiae, especially by the use of new quantitative real-time polymerase chain reaction (PCR) methods. Moreover, recent breakthroughs have occurred leading to the development of new axenic culture for intracellular bacteria including Tropheryma whipplei and Coxiella burnetii that will open the way in the future for development of new axenic media for these intracellular bacteria to facilitate antibiotic susceptibility testing (8). Resistance to antimicrobials has been infrequent among the organisms considered in this chapter, although resistance of genital mycoplasmas to tetracycline has been reported (9), as well as resistance of Chlamydia trachomatis to erythromycin and tetracycline (10,11).

Chlamydiae are obligate intracellular bacteria that undergo a complex growth cycle. Three human pathogens, C. trachomatis (4), Chlamydia psittaci (12), and Chlamydia pneumoniae (TWAR) (13), occur in this genus of obligate, intracellular parasites. C. trachomatis is a major human pathogen and is probably the most prevalent sexually transmitted pathogen in the United States (14), C. psittaci is a mainly animal pathogen that occasionally causes pneumonia in humans (15), and C. pneumoniae is an important cause of community-acquired respiratory infections and is responsible for an average of 10% of cases of pneumonia and 5% of cases of bronchitis and sinusitis. Although many research groups perform antimicrobial susceptibility testing of Chlamydia organisms, there is not a standardized methodology or a uniformly accepted interpretation of results. The techniques used for susceptibility testing of these organisms is similar and involve inoculating cell monolayers with the bacteria and incubating them in the presence of serial dilutions of antibiotic (16). However, detection of bacteria varied from enumeration of bacterial inclusions after staining (Giemsa or immunofluorescence) to quantification using reverse transcriptase-polymerase chain reaction (RT-PCR) (17) or flow cytometry (18). Because of the importance of these pathogens in human disease, susceptibility testing of Chlamydia has been extensive (Table 7.1)

Cell Lines and Organism

A number of different cell lines have been used to propagate C. trachomatis, including McCoy, HeLa 229, and BHK-21 (clone 13). According to a recent study, it seems that the recommended cell line for susceptibility testing for C. trachomatis should be the McCoy line, and Hep-2 should be used for C. pneumoniae. To render these cells more susceptible to infection, a variety of treatments have been employed, including cycloheximide, diethylaminoethyl (DEAE)-dextran, 5-iodo-2-deoxyuridine, cytochalasin B, and irradiation. However, there is general agreement that the treatment of choice is cycloheximide. Cycloheximide is used at a concentration of 0.5 to 2.0 g/mL. Each lot should be tested for potency by dose-response curve analysis because this varies, and the optimum concentration for any lot can be determined only by experimentation. An acceptable growth medium is Minimum Essential Eagle Medium (EMEM) supplemented with 2 mmol/L glutamine, 4.4% (wt/vol) sodium bicarbonate, and 10% (vol/vol) fetal bovine serum (19).

McCoy cells should undergo at least two passages in an antibiotic-free medium to ensure that all traces of antibiotic in the growth medium have been removed. Some investigators carry out 10 to 15 passages in antibiotic-free medium before using the cells for susceptibility testing (4,19). When microdilution plates are used, the cells are seeded at a concentration of 3 × 10⁵ cells per well, and the plates are incubated for 48 to 72 hours, at the end of which time the cells should have formed a subconfluent monolayer (4).

Despite the widespread use of HeLa and McCoy cells for susceptibility studies with C. trachomatis, these cells may not provide a relevant in vitro environment for such testing. This was demonstrated in a publication by Wyrick et al. (20), who showed that the minimal inhibitory concentration (MIC) for azithromycin was substantially lower with polarized human endometrial gland epithelial cells than with similar nonpolarized cells (0.125 g/mL and 0.5 g/mL, respectively). This was later confirmed by Paul et al. (21). However, the use of such cells is still not practical for most laboratories, and further work will have to be done to determine whether the extra effort of employing such cell systems is going to yield results that are more clinically relevant.

Chlamydial organisms should undergo at least one passage in antibiotic-free tissue culture cells, and sufficiently high-titered pools should be developed so that 10² to 10³ inclusion-forming units (IFU) per coverslip are achieved. Because relatively little variation in susceptibility has been seen among different clinical isolates, investigators either have used well-characterized laboratory isolates (19) or recent clinical isolates (22–25). The latter are preferable if an attempt is being made to detect whether resistance is developing in current clinical isolates.

Antimicrobial Susceptibility Testing

Some workers prefer 1-dram shell vials, as opposed to microdilution plates, because the larger surface area of the coverslips (diameters vary from 10 to 12 mm) used with the vials makes it somewhat easier to detect low numbers of inclusions. The larger area provides greater assurance that a valid end point (MIC) will be obtained. On the other hand, if one uses high-titered pools of chlamydiae so that 10² to 10³IFU per coverslip are achieved, there is no reason why microdilution plates with 96 wells cannot be used (4,26). An alternative is to use 24-well plates with wells of a 13-mm diameter, which effectively circumvents the problem of too small a surface area (27). Probably, perfectly valid results can be obtained with microdilution plates, and if larger numbers of clinical isolates are to be tested or if larger numbers of compounds are to be evaluated, then the use of microdilution plates (or plates with at least 24 wells) is the only practical alternative.

Recommended Technique: Microdilution Plate Method

Prior to inoculation, the monolayers are exposed to DEAE-dextran at a concentration of 30 g/mL for 10 to 30 minutes (4,28). Each of the 24-well plates is inoculated with an inoculum of chlamydiae that yields 5 × 10³ IFU/mL. The infectious inoculum (0.1 mL) is centrifuged onto the monolayer at 1,200 g for 60 minutes at room temperature. This centrifugation step is essential to infect cells with all C. trachomatis strains other than those that cause lymphogranuloma venereum. After centrifugation, the growth medium is removed and replaced with EMEM supplemented with glucose (5 mg/L), cycloheximide (generally 1 mg/L), 3% fetal bovine serum, and serial twofold dilutions of each antibiotic to be tested (19). All tests are performed in triplicate. The plates are then incubated at 35°C in a CO₂incubator for 48 to 72 hours. Then the coverslips are removed from the wells, fixed in absolute methanol-acetone, and stained (4,28).

Detection of Inclusions

Most investigators who have done susceptibility testing of C. trachomatis have used iodine staining for the detection of inclusions (6). One of the major disadvantages of this stain is that the inclusions may not be detected if they are particularly small or aberrant in shape. Aberrantly shaped inclusions are particularly common when C. trachomatis is cultured in the presence of β-lactam antibiotics (29). Giemsa staining has also been used, but it is evident that either direct or indirect fluorescent staining of the monolayers is the most sensitive method for detecting chlamydial antigen in cell monolayers (4,19,29). Initially, fluorescent staining for susceptibility testing used an indirect fluorescent antibody technique employing a polyclonal antibody raised in a rabbit against the same serovar E strain used in the susceptibility studies (30). However, with the availability of fluorescein-conjugated monoclonal antibodies (Syva, Palo Alto, CA; Ortho Clinical Diagnostics, Raritan, NJ; or Kallestad, Chaska, MN) to C. trachomatis, direct fluorescent staining has become the method of choice for detecting chlamydial antigen in tissue culture (4,22–24,26,31–37). In using fluorescent stains, the manufacturers’ directions should be followed. When Giemsa staining and direct immunofluorescent (using the Syva monoclonal antibody) techniques were compared, it was evident that the MICs obtainable with the monoclonal antibody were about two times higher than those obtained with Giemsa staining (29). Enzyme-linked immunosorbent assays (ELISA) have also been used to detect chlamydial antigen, and these yield MIC values comparable to those obtained with the immunofluorescent method (38). A commercially available enzyme immunoassay (Chlamydiazyme; Abbott Laboratories, North Chicago, IL) has also been used to detect antigen, and the results were similar to those obtained using a genus-specific monoclonal antibody (Ortho Clinical Diagnostics) in an immunoperoxidase test (39).

The MIC is defined as the lowest concentration of antibiotic that completely inhibits inclusion formation after 48 to 72 hours of incubation, and the minimal bactericidal concentration (MBC) is defined as the lowest concentration of antibiotic that completely inhibits the development of inclusions when the cells are disrupted at 48 to 72 hours and passed into tissue culture medium that is free of antibiotics.

Other Assays

Antibiotic susceptibility testing for Chlamydia has also been performed using flow cytometry (18). In this assay, evaluation of antibiotic activity was done at the 25-hour time point, and cells were best permeabilized using the Ortho/Permeafix treatment. The mean fluorescence intensity (MFI) of cells was determined by this method after staining of chlamydial inclusions with an anti-Chlamydia fluorescent monoclonal antibody. Calculation of the inhibitory concentration 50 (IC₅₀), defined as the antibiotic concentration required to reduce the drug-free control MFI by 50%, by flow cytometry allowed a more objective and precise evaluation of antibiotic activity than MIC (18).

Finally, an RT-PCR–based method has been developed for antimicrobial susceptibility testing of C. pneumoniae (40) and C. trachomatis (17). The results obtained in these studies were in the range previously reported using immunofluorescent staining, and the MICs obtained by RT-PCR were consistently higher (17,40). The advantage of the RT-PCR technique over ordinary PCR methods is that only viable organisms will produce RNA.

Results of Susceptibility Testing

Table 7.1 presents the results of MICs of antimicrobial agents against C. trachomatis and C. pneumoniae indicating the intense interest in antimicrobials with activity against these important human pathogens. In Table 7.1, as in all subsequent tables, a range is given for the MIC values, except in those instances when so few isolates were studied that only a single MIC value is available. Aminoglycosides are without any activity and can therefore be incorporated into tissue culture media used for the isolation of this organism. β-Lactamine compounds, chloramphenicol, clindamycin, imipenem, metronidazole, and vancomycin are not active against Chlamydia. The β-lactams result in the formation of aberrant inclusions but lack significant activity. Susceptibility of C. pneumoniae is similar to that of C. trachomatis, except that C. pneumoniae is resistant to sulfonamides. The most active agents against Chlamydia are the macrolides, tetracyclines, rifampin, and fluoroquinolones. However, rifampin is not used clinically because resistance develops rapidly in vitro (41).

Antimicrobial Resistance

The development of significant resistance to currently used antimicrobials has not been a problem in human isolates, although relative resistance to sulfonamides, erythromycin, rifamycins, and fluoroquinolones has been reported for C. trachomatis (10,11,42,43), arguing for the continued surveillance of current clinical isolates. One potential explanation for the lack of resistance by chlamydiae is their unique life cycle (5). However, resistance to tetracycline in swine Chlamydia suis has been reported in the Midwestern United States (44,45). Despite the lack of evidence for frequent resistance in chlamydiae in human isolates, it is clearly possible to induce resistance in the laboratory by serial passages of organisms in subinhibitory concentrations of antimicrobials (46).

Disease in humans is associated with at least four Mycoplasma species: respiratory infections with Mycoplasma pneumoniae and urogenital infections with Mycoplasma hominis, Mycoplasma genitalium, and Ureaplasma urealyticum. Finally, Mycoplasma fermentans has been isolated from patients with AIDS, and there has been conjecture about the possible role of these mycoplasmas as cofactors in the disease caused by HIV. M. fermentans has also been detected in some cases of fatal respiratory distress in immunocompetent adults (47).

The main structural characteristic of mycoplasmas is their lack of a cell wall, which makes them naturally resistant to β-lactams and all antibiotics that target the cell wall, including glycopeptides and polymyxins and agents that interfere with the synthesis of folic acid (9).

The techniques for isolating and identifying these agents are well known (48), but the techniques for performing antimicrobial susceptibility studies are less well defined (1). Both the agar dilution (49) and broth dilution (50) methods have their proponents. The two methods may yield quite disparate results for certain antibiotics, and so it is always important to consider the method used when evaluating the results of susceptibility studies (51).

A particular problem with each is that there is some drift of end points with time (a progressive increase in MIC values with prolonged incubation), as long incubation times are required for most mycoplasmas because of their slow growth. One of the probable reasons for the phenomenon of drift is that antibiotic inactivation occurs during incubation.

One advantage of the agar method is that in a mixture of sensitive and resistant strains, the two types of strains can be differentiated. With the broth method, differentiation is impossible without the cloning of isolates (49). On the other hand, from a purely clinical standpoint, it is probably of little importance to detect a mixture of resistant and susceptible strains, although it could be of considerable research interest (52).

Tanner et al. (53) adapted commercially available Sensititre broth microdilution plates for the susceptibility testing of Mycoplasma hyopneumoniae, and this technique was also successfully employed by Poulin et al. (54) for testing AIDS-associated mycoplasmas. The technique yields results comparable to those obtained with the macrodilution method.

Limb et al. (55) have utilized the measurement of ATP bioluminescence for the susceptibility testing of mycoplasmas. Using this technique, they were able to demonstrate good correlation with conventional methods and could achieve results within 6 hours.

Media and Organisms

Actively growing broth cultures are frozen at −70°C. A suitable medium for M. pneumoniae, M. fermentans, Mycoplasma incognitus, and M. genitalium is SP-4 medium (56), and a suitable medium for both of the genital mycoplasmas is 10-B broth. 10-B broth is usually made with penicillin, which should be omitted in susceptibility testing. Commercially prepared 10-B broth, as well as other specialized media for the isolation and propagation of mycoplasmas, may be obtained from commercial sources (e.g., Regional Media Laboratories, Lenexa, KS) and can be ordered without antibiotics for susceptibility testing.

For the culture of M. hominis, arginine, rather than urea, is incorporated into the medium (48). An aliquot of the culture is thawed, and serial dilutions are carried out to determine how many color changing units (CCUs) are present per milliliter. One CCU is the minimum inoculum required to produce enough growth to cause a color change in the phenol red indicator.

Antimicrobial Susceptibility Testing

The authors favor the broth dilution technique because it can be carried out in microdilution plates, allowing relatively large numbers of isolates to be tested against a reasonable number of different antimicrobials (57). Furthermore, the technique is adaptable for a variety of mycoplasmas, and when tests are carried out in triplicate, very good reproducibility is noted (57,58).

In the case of U. urealyticum, there has been good agreement noted between the more laborious tube dilution technique and the microdilution technique (59). Furthermore, both the MIC and the MBC can be determined using the broth dilution technique, whereas only the MIC can be determined using the agar dilution technique. Finally, the antibiotic broth dilution technique is really just an adaptation of the metabolic inhibition test (48), which has been used for identifying and serotyping mycoplasmas as well as for serodiagnosis. For this reason, laboratories may already be familiar with the basic components of the broth dilution technique.

The metabolic inhibition technique depends on the presence of either antibodies or, in the case of susceptibility testing, antimicrobials inhibiting the growth of the mycoplasmas. Inhibition of growth is detected by the lack of color change of a pH indicator, generally phenol red. Suitable substrates are included in the growth medium for the varying mycoplasmas: glucose in the case of M. pneumoniae, arginine for M. hominis, and urea for U. urealyticum.

Recommended Technique: Broth Dilution Technique

The broth dilution method is performed in 96-well plates with a volume of 200 µL in each well. Each stock antibiotic is added in a volume of 0.025 mL to a well of a microdilution plate, generally in triplicate (1,57). An aliquot of a previously frozen (−70°C), actively growing broth culture is thawed on the day of the assay and added to 50 mL of the appropriate broth medium (SP-4 for M. pneumoniae and 10-B broth for M. hominis and U. urealyticum) for each antibiotic to be tested. The stock culture is diluted to yield 10³ to 10⁴ organisms per microdilution well. To further establish how many CCUs have been added to each well, 10-fold dilutions of the inoculum are made to verify that at least 10³ CCUs but no more than 10⁵ CCUs have been added to each well. Inoculated broths are incubated for 2 hours at 37°C before these broths are added to the microdilution plates. Mycoplasma suspensions are added in 0.175-mL aliquots to each well containing antibiotics. The plates are sealed in plastic bags containing sterile gauze moistened with distilled water and are incubated at 35°C to 37°C under atmospheric conditions.

Three controls are included: (a) a broth control with no mycoplasmas, (b) a drug control consisting of the maximum drug concentration tested in broth alone, and (c) a mycoplasma control consisting of the mycoplasma suspension alone in a total of 0.2 mL of broth. Plates are examined after 17 to 20 hours of incubation and once daily until growth is noted in the mycoplasma control well. The MIC will be generally available for U. urealyticum at 24 hours, for M. hominis at 48 hours, and for M. pneumoniae after 5 or more days.

Determination of Minimal Inhibitory Concentration and Minimal Bactericidal Concentration

With SP-4 medium and 10-B broth, growth of U. urealyticum sufficient for determination of the MIC occurs overnight. Comparable times are 24 to 48 hours for M. hominis and 3 to 5 days for M. pneumoniae. Often, investigators determine both initial and final MICs (57); the initial MIC is the minimum amount of antibiotic required to inhibit any color change of the broth when the control well (containing organisms but no antibiotic) first shows a color change, and the final MIC is the minimum concentration of antibiotic that prevents a color change over a period of 2 consecutive days. The final MIC is employed with mycoplasmas because of their slow growth characteristics, which result in the drift of the MIC. In fact, the final MIC may be as much as eight times higher than the initial MIC for some antimicrobials (57).

It is also possible to determine the MBC, by diluting the broth from wells showing no color change in antibiotic-free medium. Generally, this is a 20-fold dilution, which is usually sufficient to dilute the antibiotic to a level below the antibiotic’s MIC value (1), but hopefully not to a point where organisms can no longer be detected. An alternate method avoids these possible pitfalls by filtering the broth from wells with no color change through a filter with a pore size of 220 nm, washing the filter free of residual antibiotic, and culturing the filter (60). Using the latter technique, Taylor-Robinson and Furr (60) were able to show that the macrolide rosaramicin acted in a purely mycoplasmastatic fashion on some ureaplasmas.

In the case of ureaplasmas, there is a self-sterilizing effect, so that by 24 hours, there is often a precipitous fall in the number of organisms (1). Because Taylor-Robinson and Furr (60) showed that there is no change in the MIC values for these organisms between 5 and 25 hours, subculturing for the determination of MBC values can be done as early as 5 hours, which avoids any problems that may be caused by the self-sterilizing phenomenon (1).

Results of Susceptibility Testing

The results of susceptibility testing on mycoplasmas are summarized in Tables 7.2 and 7.3. Mycoplasmas lack peptidoglycan and penicillin-binding proteins and thus are naturally resistant to β-lactam antibiotics. Moreover, they are also resistant to rifampin, owing to the particular structure of their RNA polymerase, and also to polymyxins, nalidixic acid, sulfonamides, and trimethoprim. Tetracyclines, erythromycin, clindamycin, chloramphenicol, aminoglycosides, and fluoroquinolones have been shown to have activity against one or more mycoplasmal species (61).

Mycoplasma pneumoniae

Erythromycin and tetracyclines are usually active against M. pneumoniae. Resistance to erythromycin has now been described (62,63), although resistance to tetracycline has not been documented (64). Azithromycin and telithromycin are more active against M. pneumoniae in vitro than erythromycin, clarithromycin, or roxithromycin (32,65,66). Fluoroquinolone compounds are also active against M. pneumoniae, but in vitro studies have shown that they are not as effective as macrolides (67–69). Clinical strains with acquired resistance to macrolides have recently emerged worldwide with resistances to erythromycin and azithromycin mainly due to mutations in 23S rRNA (64).

Mycoplasma hominis

M. hominis is usually naturally susceptible to tetracyclines but tetracycline-resistant isolates containing DNA sequences homologous to the streptococcal determinant tetM have been reported (52). It has been convincingly demonstrated that tetM is not a plasmid and that it is present in both species of genital mycoplasmas. Resistance to tetracycline in vitro is associated with failure of tetracycline treatment to eradicate M. hominis (9). A recent study from Germany has shown that the prevalence of resistance to tetracyclines and fluoroquinolones has increased between 1989 and 2004, but doxycycline still remains the drug of choice for the treatment (70). The new glycylcyclines have been shown to be active in vitro against M. hominis strains resistant to other tetracyclines. Because of this emerging resistance, it is proving to be increasingly difficult to devise suitable antimicrobial regimens for the effective therapy of genital mycoplasma infections (9,71–73). Clindamycin can be used for the treatment of M. hominisinfections resistant to tetracyclines, and erythromycin or quinolones can be used for tetracycline-resistant U. urealyticuminfections (9,39). Usually, fluoroquinolone compounds are active against M. hominis, but resistance to fluoroquinolones has been reported and is associated with mutations in DNA gyrase (74). A fatal case of a disseminated infection with clindamycin and ciprofloxacin-resistant M. hominis has been recently reported in Germany (75). M. hominis is resistant to erythromycin, roxithromycin, azithromycin, and clarithromycin but remains susceptible to josamycin (Table 7.3).

Ureaplasma urealyticum

Tetracyclines and fluoroquinolone compounds are active against U. urealyticum. Erythromycin is generally active against ureaplasmas, but resistance has been noted (14) and is associated with specific mutations in the 23S rRNA gene (76).

All members of the genus Rickettsia are obligate, gram-negative, intracellular bacteria. The genus comprises typhus group rickettsiae, which include Rickettsia prowazekii, the agent of epidemic typhus, and Rickettsia typhi, the agent of murine typhus; Orientia tsutsugamushi, the agent of scrub typhus (77); and spotted fever group (SFG) rickettsiae. The number of recognized SFG rickettsioses has recently increased. The six SFG rickettsioses previously described are Rocky Mountain spotted fever, caused by Rickettsia rickettsii; Mediterranean spotted fever, caused by Rickettsia conorii subsp conorii; Siberian tick typhus, caused by Rickettsia sibirica subsp sibirica; Israeli spotted fever, caused by R. conorii subsp israelensis (77); Queensland tick typhus, caused by Rickettsia australis; and rickettsialpox, caused by Rickettsia akari. Since 1984, 12 new SFG rickettsiosis have been described (77): the Japanese or Oriental spotted fever, caused by Rickettsia japonica and described in 1984; Flinders Island spotted fever, caused by Rickettsia honei and described in 1991; Astrakhan fever, caused by R. conorii subsp caspia and reported in 1991; African tick-bite fever, caused by Rickettsia africae and described in 1992; a new spotted fever due to Rickettsia mongolitimonae, reported in France in 1996; Rickettsia slovaca infection, reported in 1997; Rickettsia helveticainfection, described in 2000; flea-borne rickettsioses, caused by Rickettsia felis and reported in 2001; Rickettsia aeschlimannii infection, reported in 2001; Rickettsia heilongjiangensis infection; Rickettsia parkeri infection, reported in 2003; Rickettsia massiliae; and Rickettsia marmionii (77).

All of the members of the genus Rickettsia are obligate intracellular pathogens and, therefore, require either animal models, embryonated eggs, or tissue culture for susceptibility assays (78). It would appear that, of the in vitro techniques now available, the plaque assay and a colorimetric assay (78) are the most practical for evaluating antiinfectives for this group of organisms. However, because of the technical difficulties in working with these agents, susceptibility testing will probably be confined to relatively few laboratories. Furthermore, it is evident that when in vivo techniques, such as suppression of lethality in chicken embryos, are compared with in vitro techniques, such as the plaque assay, the results may be somewhat discrepant (79). The two assays (plaque assay and colorimetric assay) depend on the induction of cytopathic effects and plaque formation in cell cultures by the rickettsiae, but some rickettsiae do not normally cause cytopathic effects in primary cultures (80,81). Recently, Ives et al. (82,83) described a new assay that uses immunofluorescent staining, which avoids the problem of a lack of cytopathic effects. Very recently, we have developed a new quantitative PCR DNA assay using the LightCycler system for the evaluation of antibiotic susceptibilities of three rickettsial species, including R. felis, a rickettsial species that does not induce plaque in cell cultures (81).

Cell Lines and Organisms

Organisms used for these studies are laboratory strains. For example, in the case of R. rickettsii, the Sheila Smith strain is used (78), and for R. conorii, the American Type Culture Collection VR 141 Moroccan strain is used (84). Only a few studies on in vitro antibiotic susceptibilities of SFG rickettsiae other than R. conorii, R. rickettsii, and R. akari are available. We recently reported an extensive study that investigated the reaction of 27 rickettsiae to 13 antimicrobials (Table 7.4) (80).

Antimicrobial Susceptibility Testing

Reference Method: Plaque Assay

Vero cell monolayers seeded 24 hours before use in round, plastic, tissue culture Petri dishes (60 mm; Corning Glass Works, Corning, NY) are infected with 1 mL of a solution containing 4 × 10³ plaque-forming units (PFU) of the desired rickettsial strain (78). After 1 hour of incubation at room temperature (22°C), the plates are overlaid with 4 mL of a medium containing Minimum Essential Eagle Medium (Gibco Laboratories, Grand Island, NY), 2% newborn calf serum, 2% (N-[2-hydroxyethyl]) piperazine-N ′-[2-ethanesulfonic acid]) (HEPES, Sigma-Aldrich, St. Louis, MO), and 0.5% agar. The antibiotic solutions are added to the medium to obtain the desired final concentrations (an antibiotic-free control plate is also included), and the plates are incubated for 4 to 7 days at 35°C in a CO₂ incubator. All antibiotics are assayed in triplicate, at a minimum. After incubation, the monolayers are fixed with 4% formaldehyde and stained with 1% crystal violet in 20% ethanol. The MIC is the lowest concentration of the agent tested causing complete inhibition of plaque formation, compared with the drug-free controls. Plates may then be photographed to obtain a permanent record of the results. The major problem with this technique is that some rickettsial strains may not induce the formation of plaques in cell cultures. In this case, several passages in various cell lines may allow selection of variants able to produce plaques.

Rickettsiacidal activity can be determined from this assay by staining surviving cells with either Gimenez or immunofluorescent stain (79). The minimal concentration of antibiotic that completely sterilizes the monolayer is defined as the minimum rickettsiacidal concentration.

Dye-Uptake Assay

Flat-bottomed microdilution plates are seeded with 1.5 × 10⁴ Vero cells (suspended in a solution of EMEM, 5% newborn calf serum, and 2 mmol/L L-glutamine) per well and subsequently infected with varying concentrations of a suspension of Rickettsia organisms (78). The Vero cell suspension (100 µL) is added to each well. The infectious inoculum is added to individual wells in a final volume of 50 µL. For each 96-well plate, the first horizontal row of 8 wells contains no rickettsiae, the second horizontal row is inoculated with 2,000 PFU, the third horizontal row with 200 PFU, and the fourth with 20 PFU (Fig. 7.1). Two antibiotics are tested per plate, four concentrations of each antibiotic are tested, and each concentration of antibiotic is replicated 12 times; that is, each row contains the same antibiotic at the same concentration. Antibiotics are added in 50-µL volumes. Incubation is then carried out at 36°C for 4 days in a CO₂ incubator.

After this, the medium is removed, 50 µL of neutral red dye (0.15% in saline, pH 5.5; Sigma Chemical Co, St. Louis, MO) is added to each well, and the plate is incubated for 60 minutes at 36°C. Unincorporated dye is then washed (three washes) from the cells using phosphate-buffered saline (pH 6.5). Incorporated dye is removed from the well using 100 µL of phosphate-ethanol buffer (10% ethanol in phosphate-buffered saline, adjusted to pH 4.2).

Finally, the optical density (OD) of the solution is read at 492 nm with a multichannel spectrophotometer designed for use with microdilution plates (EIA Autoreader, model EL310; BioTek Instruments, Winooski, VT). The mean OD of the control wells (containing only Vero cells) is assigned a value of 1, and the mean OD of the wells containing 2,000 PFU is assigned a value of 0 (Fig. 7.1). The MIC is considered to be any OD value that falls between the mean OD of the wells containing 20 PFU and the mean OD of the control wells containing only Vero cells.

This assay is an adaptation of an assay used to determine the efficacy of agents against herpes viruses (85) and is dependent on the fact that intact cells take up neutral red dye. Hence, wells that contain fewer cells take up less neutral red and yield lower OD values. This test is, therefore, a derivative of the plaque assay but has the advantage of using microdilution technology as well as an automated means of reading the plates.

Other Assays

Immunofluorescence Assay.More recently, an immunofluorescence assay was described by Ives (83). In this model, Vero cells cultured in wells of chamber culture microscope slides were infected with rickettsiae. After incubation of cultures for 3 hours at 37°C in a 5% CO₂ atmosphere, cell supernatants were replaced by new medium containing various concentrations of the antibiotics to be tested. Drug-free cultures served as controls. Cell culture monolayers were then fixed with methanol and stained using an immunofluorescence assay to reveal the presence of immunofluorescent foci (clusters of rickettsiae) in 25 random fields for each well. The minimal antibiotic concentration allowing complete inhibition of foci formation as compared with the drug-free controls was recorded as the MIC.

LightCycler Polymerase Chain Reaction Assay.Cells cultured in 24-well plates were infected with rickettsiae and incubated for 7 days at 37°C in a 5% CO₂ atmosphere with medium containing various concentrations of the antibiotics to be tested. Wells were harvested each day for 7 days and stored at −20°C before the PCR assay. Real-time PCR was performed on LightCycler instrumentation (Roche Biochemicals, Mannheim, Germany) (86). The specificity of amplification can be confirmed by melting curve analysis. Single melting peaks can be generated by depicting the negative derivative of fluorescence versus temperature (−dF/dT) over the course of a gradual PCR product melt.

Extraction of DNA.After thawing, harvested tubes were centrifuged at 5,000 rpm for 10 minutes, supernatant was discarded, and pellet was washed twice with sterile distilled water and finally resuspended with 200 µL of sterile distilled water. Extraction of the DNA was performed using Chelex (biotechnology-grade chelating resin, Chelex 100, Bio-Rad, Richmond, CA) at 20% in sterile water. Briefly, 500 µL of Chelex was added to each tube, then the tubes were vortex-mixed and placed in a boiling water bath for 30 minutes. The tubes were then centrifuged at 14,000 rpm for 10 minutes, and supernatant was harvested and stored in sterile tubes at 4°C before use.

Polymerase Chain Reaction Master Mix.Master mixes were prepared by following the manufacturer’s instructions, using the primers CS877F (5′-GGG GGC CTG CTC ACG GCG G-3′) and CS1258R (5′-ATT GCA AAA AGT ACA GTG AAC A-3′) of the citrate synthase gene. The 20-µL sample volume in each glass capillary contained the following: for all single experiments, 2 µL of LightCycler DNA Master SYBR Green (Roche Biochemicals, Mannheim, Germany), 2.4 µL of MgCl₂ at 4 mM, 1 µL of each primer at 0.5 µM, 11.6 µL of sterile distilled water, and 2 µL of DNA.

Polymerase Chain Reaction Cycling and Melting Curve Conditions.After one-pulse centrifugation to allow mixing and to drive the mix into the distal end of each tube, glass capillaries were placed in the LightCycler instrument. The amplification program included an initial denaturation step consisting of 1 cycle at 95°C for 120 seconds and 40 cycles of denaturation at 95°C for 15 seconds, annealing at 54°C for 8 seconds, and extension at 72°C for 15 seconds, with fluorescence acquisition at 54°C in single mode. Melting curve analysis was done at 45°C to 90°C (temperature transition, 20°C per second), with stepwise fluorescence acquisition by real-time measurement of fluorescence directly in the clear glass capillary tubes. Sequence-specific standard curves were generated using 10-fold serial dilutions (10⁵to 10⁶ copies) of a standard bacterial concentration of Rickettsia organisms. The number of copies of each sample transcript was then calculated from a standard curve using the LightCycler software. The MIC was defined as the first antibiotic concentration allowing the inhibition of growth of bacteria as compared with the number of DNA copies at day 0. Experiments were made twice in duplicate. This technique is specific, reproducible, easy to perform, and rapid. It can also be used to measure the number of DNA copies at any time, and we were able to perform for the first time a kinetic of the growth of Rickettsiaorganisms even if the bacteria did not lead to plaque in vitro in cell cultures.

Results of Susceptibility Testing

The results of susceptibility testing of Rickettsia species are presented in Table 7.4. Sensitivities to amoxicillin (MICs from 128 to 256 µg/mL), gentamicin (MICs from 4 to 16 µg/mL), and co-trimoxazole were poor (80,81). Doxycycline was the most effective antibiotic against all strains tested, with MICs ranging from 0.06 to 0.25 µg/mL. The MICs of thiamphenicol ranged from 0.5 to 4 µg/mL, and the MICs for fluoroquinolone compounds ranged from 0.25 to 2 µg/mL. Among the macrolide compounds, josamycin was the most effective antibiotic, with MICs ranging from 0.5 to 1 µg/mL. Typhus group rickettsiae were susceptible to erythromycin (MICs from 0.125 to 0.5 µg/mL), whereas SFG rickettsiae were not (MICs from 2 to 8 µg/mL), and this difference is likely due to mutations in L22 ribosomal protein for SFG rickettsiae (87). Recently, we demonstrated that the new ketolide compound telithromycin was very effective against typhus group rickettsiae and SFG rickettsiae, with MICs ranging from 0.5 to 1 µg/mL (97). Susceptibilities to rifampin varied: typhus group rickettsiae and most SFG rickettsiae were susceptible (MICs from 0.03 to 1 µg/mL), but a cluster including R. massiliae, Rickettsia montana, Rickettsia rhipicephali, R. aeschlimannii, and strain Bar 29 were more resistant (MICs from 2 to 4 µg/mL). This relative resistance to rifampin was linked to natural mutations in the rpoB gene (88).

Ehrlichioses are emerging infectious diseases caused by obligate, gram-negative, intracellular bacteria belonging to the Proteobacteria α subgroup (89). The genus Ehrlichia is divided into three genogroups: the group Neorickettsia, with Neorickettsia sennetsu, Neorickettsia risticii, and Neorickettsia helminthoeca; the group Ehrlichia, with Ehrlichia canis, Ehrlichia chaffeensis, Ehrlichia rumitantium, Ehrlichia muris, and Ehrlichia ewingii; and the group Anaplasma with Anaplasma platys, Anaplasma marginale, and Anaplasma phagocytophilum (89,90).

They are responsible for human and animal diseases. E. chaffeensis is the agent of human monocytic ehrlichiosis (HME); A. phagocytophilum, the agent of human granulocytic ehrlichiosis (HGE); and E. canis, the agent of canine ehrlichiosis. In vitro and in vivo antibiotic susceptibility studies have been carried out on various species of Ehrlichia. All have found that doxycycline and rifampin are highly effective against ehrlichiae, and thus they are currently preferred for treating animal and in human ehrlichiosis.

Antimicrobial Susceptibility Testing

Animal Models

N. sennetsu was first isolated in mice, and subsequently, infections in mice have been used as a model for Sennetsu fever. Although the growth of N. sennetsu in mice is much slower than the growth of other rickettsiae, treatment of mice with cyclophosphamide prior to inoculation has been found to enhance the growth of N. sennetsu (91), and this technique has been used for the preparation of antigen in mice. The first study of antibiotic susceptibility in mice for N. sennetsu has shown that erythromycin, sulfisoxazole, penicillin, streptomycin, polymyxin B, bacitracin, and chloramphenicol were ineffective even at high concentrations (92). Chlortetracycline was more effective than oxytetracycline and tetracycline. Further studies have evaluated the effect of tetracycline therapy on spleen size as a percentage of body weight and on the splenic infectious burden in mice infected with N. sennetsu (93,94). In mice, in which tetracycline therapy was initiated at the same time as inoculation, there were no detectable ehrlichiae in the spleen (93). Therefore, it would appear that the time of initiation of treatment may be important in controlling the course of infection with N. sennetsu and that delayed therapy may allow the development of chronic infections.

Cell Culture Model

The susceptibility of Ehrlichia species to various antibiotics has been tested using ehrlichiae-infected contact-inhibition-growth cell lines incubated for 48 to 72 hours in the antibiotic concerned. Thereafter, the antibiotic-containing media is removed, and ehrlichiae-infected cells are incubated with antibiotic-free media for at least 3 more days. The number of ehrlichiae-infected cells is counted every day, and an antibiotic is considered ineffective if the number of ehrlichiae-infected cells after exposure to the antibiotic is similar to that of noninfected control cells. If the number of ehrlichiae-infected cells is found to decrease during incubation with an antibiotic, the antibiotic is regarded as being bactericidal. Antibiotics are considered bacteriostatic if there is no increase or decrease in ehrlichiae-infected cells when the antibiotic is present but the number of infected cells increases when antibiotic-free media is provided.

Results of Susceptibility Testing

Results of susceptibility testing for A. phagocytophilum, E. canis, and E. chaffeensis using the Diff-Quick assay are presented in Table 7.5. The in vitro susceptibility of N. sennetsu Miyayama strain to eight antibiotics was determined using Diff-Quick staining of infected P388D1 cells over a 5-day period (95). In this study, it was also demonstrated that Diff-Quick staining was as reliable as immunofluorescence assay for detecting infected cells. It was found that N. sennetsu was not susceptible to penicillin, gentamicin, co-trimoxazole, erythromycin, and chloramphenicol, whereas rifampin, doxycycline, and ciprofloxacin were effective, with MICs of 0.5, 0.125, and 0.125 µg/mL, respectively.

The in vitro antibiotic susceptibility of E. chaffeensis was studied recently (96,97) by means of a microplate colorimetric assay using ehrlichiae-infected DH82 cell culture. The percentage of infected cells was determined each day by Diff-Quick staining, which has been found to stain only viable organisms. On the third day of incubation, the antibiotic-containing medium was removed and replaced with antibiotic-free medium. Using these methods, it was found that E. chaffeensis was sensitive to 0.5 µg/mL of doxycycline and 0.125 µg/mL of rifampin. Chloramphenicol, co-trimoxazole, erythromycin, telithromycin penicillin, gentamicin, and ciprofloxacin were not effective against E. chaffeensis.

The HGE agent is sensitive to doxycycline, ofloxacin, ciprofloxacin, and trovafloxacin but is resistant to clindamycin, co-trimoxazole, erythromycin, azithromycin, ampicillin, ceftriaxone, and imipenem (98–100). Chloramphenicol and aminoglycosides only display a poor bacteriostatic activity and are never bactericidal (99). Fluoroquinolones are more active in vitro against A. phagocytophilum than against E. chaffeensis and E. canis (Table 7.5).

Fluoroquinolones might represent a potential therapeutic alternative to tetracycline for HGE, but they have not received U.S. Food and Drug Administration (FDA) approval for use in children and pregnant women. Moreover, a Gyr A–mediated resistance in the related species E. canis and E. chaffeensis has recently been described and can explain this difference (101).

Recently, an evaluation of antibiotic susceptibilities against E. canis, E. chaffeensis, and A. phagocytophilum has been performed using a new real-time PCR assay (102). Although doxycycline and rifampin were highly active against the three species, there was a heterogeneity of susceptibility for fluoroquinolones. Interestingly, macrolide compounds and telithromycin were not effective because of numerous point mutations in their 23S RNA genes (102). A new real-time PCR assay for antibiotic susceptibility testing has been also developed and tested for 18 O. tsutsugamushi fresh isolates that demonstrates an intrinsic heterogeneity of susceptibility against fluoroquinolone compounds due to a point mutation in their quinolone resistance-determining region (QRDR) domain (103).

C. burnetii, the agent of Q fever, is an obligate, intracellular bacterium that multiplies within acidic vacuoles of eukaryotic cells (104). C. burnetii is classified in the family of Rickettsiaceae, where it belongs to the Proteobacteria γ subgroup based on 16S-rRNA sequence analysis (105). C. burnetti is a bioterrorism agent that is resistant to heat and drying and can survive in the environment for months. It is also highly infectious by the aerosol route. Owing to these features, Q fever has been investigated and developed as a bioweapon. If used, it would not generate mass fatalities but rather act as an incapacitating agent (106,107). Acute Q fever is the primary infection, and in specific hosts, it may become chronic (108). A few patients (~0.7%) suffer from chronic Q fever, which in most cases corresponds to chronic endocarditis, especially in patients with previous cardiac valve defects and/or with a cardiac valve prosthesis, in immunocompromised patients, and in pregnant women (105).

Antimicrobial Susceptibility Testing and Results

Antibiotic susceptibility testing of C. burnetii was difficult because this organism is an obligate, intracellular bacterium. Previously, three models of infection have been previously developed: animals, chick embryos, and cell culture. Since the previous version of this chapter, quantitative real-time PCR assay have been developed for antibiotic susceptibility testing of C. burnetii (109,110) that have led to the possibility to test new isolates and compounds, including doxycycline-resistant isolates (110,111). Moreover, an axenic medium has been recently described for the culture of C. burnetii (112,113) that will probably facilitate genetic manipulation and susceptibility testing of this bacterium in the future. The method that has been mostly used to test antibiotic susceptibility of C. burnetii is based on cell culture models. Torres and Raoult (118) have developed a shell vial assay with human embryonic lung (HEL) cells for assessment of the bacteriostatic effect of antibiotics, but quantitative real-time PCR could now be used routinely to assess susceptibility to antibiotics from clinical isolates (111).

Recommended Technique: The Shell Vial Assay

In this model, HEL fibroblast cells are grown in shell vials at 37°C in a 5% CO₂ atmosphere. Cell monolayers are infected with a C. burnetii inoculum previously determined to induce 30% to 50% infection of HEL cells after 6 days of incubation in the absence of antimicrobial agents, as revealed by an immunofluorescence technique with anti–C. burnetii polyclonal antibodies. The percentage of infected cells in antimicrobial-containing cultures is determined after the same incubation time using the same immunofluorescence procedure. MICs correspond to the minimum antimicrobial concentration allowing complete inhibition of growth, that is, 0% infected cells after the 6-day incubation period (Fig. 7.2).

Amikacin and amoxicillin were not effective; ceftriaxone and fusidic acid were inconsistently active (118), whereas co-trimoxazole, rifampin, doxycycline, tigecycline, clarithromycin, and the quinolones were bacteriostatic (114–116). There was a heterogeneity of susceptibility to erythromycin of the strains tested (117,118). C. burnetii can establish a persistent infection in several cell lines, including L929 mouse fibroblasts and J774 or P388D1 murine macrophage-like cells (119). Infected cells can be maintained in continuous cultures for months (120). Raoult et al., using P388D1 and L929 cells, showed that pefloxacin, rifampin, and doxycycline (121) as well as clarithromycin (115) were bacteriostatic against C. burnetii. A real-time quantitative PCR assay was recently used for antibiotic susceptibility testing on C. burnetii and proved to be more specific and sensitive than the shell vial assay (109,110). Moreover, this technique was recently used to evaluate susceptibility to antibiotics against 13 new isolates to demonstrate that telithromycin compound was effective (111). This has led also to the description of the first doxycycline-resistant human isolate from a German patient (111) for which whole genome sequence has been recently published (122).

An original model of killing assay has been developed by Maurin to assess the bactericidal activity of antibiotics against C. burnetii (123). The bactericidal activity of antibiotics in this technique is directly evaluated by titration of residual viable bacteria in persistently infected P388D1 cell cultures (Fig. 7.3). On the first day of the experiment, P388D1 cells infected with C. burnetii were harvested from a 150-cm² culture flask and seeded into 25-cm² flasks so that each flask received the same primary inoculum. Antibiotics were added to some of the flasks, and all the flasks, with or without antibiotics, were incubated for 24 hours at 37°C. Then cells were lysed, and 10-fold serial dilutions of cell lysates were distributed into shell vials containing uninfected HEL cells (124). After 6 days of incubation, C. burnetiiwere stained by indirect immunofluorescence in the shell vials. It was demonstrated that doxycycline, pefloxacin, and rifampin did not show any significant bactericidal activity. The lack of bactericidal activity was related to inactivation by the low pH of the phagolysosomes in which C. burnetii survives. Raoult demonstrated that the addition of a lysosomotropic alkalinizing agent, chloroquine, to antibiotics improved the activities of doxycycline and pefloxacin, which then became bactericidal (123).

There are several pathogens of major importance in this group, including the leptospires, Borrelia species (including B. burgdorferi, the etiologic agent of Lyme disease), and Treponema species (including T. pallidum, the etiologic agent of syphilis). Whereas B. burgdorferi can be grown in a cell-free system (2,125), T. pallidum requires tissue culture for propagation (126). Because of the complexities of working with T. pallidum, as well as the fact that it is rarely isolated from clinical material, susceptibility testing for T. pallidum will have to be carried out in specialized research laboratories whose personnel have both the interest and the competence to work with this fastidious organism.

Borrelia burgdorferi

Antimicrobial Susceptibility Testing

A modified Kelly bovine serum medium has been developed that will support the growth of laboratory-adapted strains of this organism as well as fresh clinical isolates (2). The in vitro susceptibility tests use macro- or microdilution methods and standard Barbour-Stoenner-Kelly (BSK) medium (127). Using this medium, a tube dilution susceptibility test has been developed that yields both MIC and MBC values. The MIC was defined by Berger et al. (2) as the minimum concentration of antibiotic that did not allow the spirochete to multiply, whereas Preac-Mursic et al. (128) defined the MIC as the minimum concentration that prevented growth altogether. The MBC was also determined by Berger et al. (2) by subculturing all tubes that showed inhibition of growth of the spirochete into tubes containing antibiotic-free Kelly bovine serum medium. The minimum antibiotic concentration that yielded no organisms on subculture was defined as the MBC. From these definitions of MIC and MBC, it would appear that what Preac-Mursic et al. (128) defined as the MIC was what Berger et al. (2) defined as the MBC. This indeed seems to be confirmed by the results obtained by the two groups. Thus, the MBCs obtained by Berger et al. (2) for penicillin G ranged from 0.08 to 2.5 units/mL and were comparable to the range of MICs (0.06 to 3.0 g/mL) obtained by Preac-Mursic et al. (128).

Tube Dilution Technique

The procedure developed by Berger et al. (2) is described here. In this technique, a serum-free Kelly medium is used. This medium is prepared as follows: 5 g of neopeptone (Difco Laboratories, Detroit, MI) is dissolved in 50 mL of boiling water, and after the solution cools to 37°C, the mixture is filtered through a no. 42 Whatman filter (Whatman Ltd., Maidstone, England). Bovine serum albumin (40 g) (fraction V, no. A2152; Sigma Chemical Co, St. Louis, MO) is dissolved in 200 mL of distilled water, filtered through a coarse filter, and then filtered through Whatman no. 42 paper. Next, the albumin and neopeptone solutions are added to sufficient distilled water to make a total of 900 mL.

The remaining ingredients of modified Kelly medium are as follows: 100 mL of CMRL 1066 medium with glutamine and without sodium bicarbonate (10; Gibco Laboratories, Grand Island, NY), 6.0 g of HEPES (Sigma Chemical Co, St. Louis, MO), 0.7 g of sodium citrate, 5.0 g of glucose, 0.8 g of sodium pyruvate, 0.4 g of N-acetylglucosamine, 2.2 g of sodium bicarbonate, and 1.25 g of yeastolate (Difco Laboratories, Detroit, MI). This solution is adjusted to pH 7.2 with 5.0 mol/L NaOH; after which, 200 mL of a warm 7% solution of gelatin is added. Finally, the medium is sterilized by passing it through a 0.2-m filter (Nalgene sterilization filter unit type LS; Nalge Co, Rochester, NY).

B. burgdorferi, including laboratory-adapted strains, readily grow in modified Kelly medium; by 72 hours, counts as high as 6 × 10⁶ spirochetes can be achieved. Fresh human isolates tend to clump when they grow, which is not as true for the laboratory-adapted strains. Incubation is carried out at 32°C to 33°C, and cell counts are carried out by performing serial 10-fold dilutions to 1:1,000 and examining 6 µL on a slide by dark-field microscopy. Slides are prepared by covering the sample gently with a coverslip ringed with petrolatum.

Susceptibility testing is carried out in 13 × 100-mm test tubes prepared with 5 mL of modified Kelly medium, 1 mL of antibiotic solution containing seven times the final desired antibiotic concentration, and 1.0 mL of spirochetes at a concentration of 7 × 10⁵/mL. The MIC is determined after 72 hours of incubation and is defined as the antibiotic concentration in which more than 90% of the spirochetes are motile (as determined by examination under dark-field microscopy, as described previously) and yet the number of spirochetes is not greater than the original inoculum. The MBC is determined by transferring 50 µL from all tubes showing no growth at 72 hours to 7.0 mL of modified Kelly medium. These tubes are subsequently examined at 11 days by removing 10 µL and viewing this under a dark-field microscope. The MBC is defined as the concentration of antibiotic that prevents growth. It should be noted, however, that the developers of this technique found that the MBC determinations were not reproducible (2).

Dever et al. (127) adapted the macrodilution method to a microdilution method using microtiter trays and demonstrated that the results obtained using this very efficient method were comparable to those obtained with the more laborious macrodilution method.

Recently, Hunfeld et al. (130) developed a new standardized colorimetric assay for the determination of susceptibility to antibiotics of several strains of Borrelia. This assay is based on color changes that result from actively metabolizing spirochetes after 72 hours of incubation. Briefly, Borrelia stock cultures were thawed, cultured in modified BSK medium at 33°C until the log phase of growth, and adjusted to 2.5 × 10⁷ organisms/mL as determined by enumeration with a Kova counting chamber (Hycor, Garden Grove, CA) combined with dark-field microscopy. Final concentrations of the lyophilized antibiotics were reconstituted by adding of 200 µL of the final inoculum suspension (5 × 10⁶ cells) in BSK containing phenol red (25 g/mL) as a growth indicator. Samples and growth controls were sealed with sterile adhesive plastic and cultured at 33°C with 5% CO₂. The presence or absence of growth was examined after 0, 24, 48, and 72 hours by kinetic measurement of indicator color shift at 562/630 nm applying a commercially available ELISA-reader (PowerWave 200, BioTek Instruments, Winooski, VT) in combination with a software-assisted calculation program (Microwin 3.0, Microtek, Overath, Germany). Colorimetric MICs of isolates were measured in triplicate by the quantification of growth achieved through the calculation of growth curves. Recently, two new techniques using fluorescent microscopy (BacLight viability staining) and dark-field microscopy have been successfully developed for antibiotic susceptibility testing of B. burgdorferi (131).

Results of Susceptibility Testing

The results of susceptibility testing for B. burgdorferi are presented in Table 7.6. B. burgdorferi strains of both European and North American origin have been tested against a variety of antimicrobials because of the increasing attention that this pathogen has recently attracted (125). It is evident that the macrolides, the tetracyclines, amoxicillin, and the third-generation cephalosporins all have good in vitro activity (127,132). Penicillin G activity is strain-dependent, with some strains being moderately resistant (133).

Treponema pallidum

Until recently, it was not possible to cultivate this organism. However, in 1981, Fieldsteel et al. (3) published a technique for the propagation of this treponeme in tissue culture. This cell culture was later confirmed (134) and has been used for drug susceptibility testing (126). Although there is no evidence of the emergence of resistance to benzylpenicillin (the treatment of choice for syphilis), clinically significant resistance to macrolides, an alternative to penicillin, has recently emerged in several developed countries due to point mutations in 23S rRNA genes (135,136). However, no documented resistance to tetracyclines has been reported to date in T. pallidum (136). The failure of penicillin therapy for syphilis has been particularly noted in patients with AIDS. However, this is not because of any demonstrated resistance of the treponeme to penicillin, but rather because the therapy probably does not completely eliminate viable treponemes from the host and eradication of the infection depends on host defenses lacking in patients with AIDS.

Antimicrobial Susceptibility Testing

It has been shown that 24- to 100-fold multiplication of T. pallidum can be obtained in tissue culture using Sf1Ep cottontail rabbit epithelial cells (obtainable from the American Type Culture Collection, Rockville, MD) under 1.5% to 3.0% oxygen at 33°C to 34°C (134,137). However, continuous in vitro culture has not yet been achieved (126).

Results of Susceptibility Testing

The results of susceptibility testing of T. pallidum are presented in Table 7.7. Using this procedure, antimicrobial susceptibilities have been determined with penicillin, tetracycline, erythromycin, spectinomycin, oral cephalosporins, and quinolones. The MICs achieved with the tissue culture system correlate quite well with the clinical and experimental results obtained with these antimicrobials, with the exception of spectinomycin. The single clinical isolate that has been shown by the method of inhibition of protein synthesis to be resistant to erythromycin (135) had an A to G transition mutation at position 2058 of the 23S rRNA gene (138).

Leptospira

Prior to 1989, the genus Leptospira was divided into two species, Leptospira interrogans, comprising all pathogenic strains, and Leptospira biflexa, comprising the strains from the environment. Within the species L. interrogans, there are approximately 200 serovars. The current classification of Leptospira is genotypic and now includes a number of genomospecies containing all serovars (139). A new species, Leptospira fainei, has been recently reported to cause infections in humans (140,141). Leptospirosis is a zoonosis acquired from a wide variety of domestic and wild animals. Although penicillin is considered the drug of choice because of its low MIC, there is concern that this antibiotic is not bactericidal for Leptospira species. Unfortunately, only limited in vitro studies with newer antimicrobials have been carried out.

Organism

Reference organisms obtained from the National Institute of Health (Tokyo, Japan) were used in a recent extensive study of antimicrobial susceptibility (142). These organisms are cultured in Korthof medium at 30°C, as described by Johnson and Harris (137).

Antimicrobial Susceptibility Testing

Tube dilution methodology may be used for determining the MIC and MBC (142). Inocula of 0.5 mL containing 0.5 × 10⁸ to 1.5 × 10⁸ organisms are put in each tube after the Leptospira species have been grown in Korthof medium for 5 days at 30°C. Inocula of 0.5 mL are added to tubes containing 4.5 mL of fresh Korthof medium with the desired concentration of antibiotics, and the tubes are incubated for 7 days at 30°C. The MIC is defined as the minimum concentration of antibiotic that inhibits all visible growth. The MBC was determined by taking 10 µL from each clear tube, subculturing this aliquot into 10 mL of fresh Korthof medium, and incubating these tubes for 3 weeks. The MBC is the minimum concentration of antibiotic that allows no growth in these subcultures, as determined by the absence of any visible growth.

Results of Susceptibility Testing

Oie et al. (142) studied the in vitro activity of 16 antibiotics against five serovar strains of the genus Leptospira. Five antibiotics (ampicillin, cefmetazole, moxalactam, ceftizoxime, and cefotaxime) yielded lower MICs than did penicillin G. Ceftizoxime and cefotaxime demonstrated the lowest MBCs and were more effective than penicillin G, streptomycin, tetracycline, ampicillin, and cefmetazole.

Because of the failure of penicillin to prevent a laboratory-acquired case of leptospirosis due to L. interrogans subgroup icterohaemorrhagiae, Broughton and Flack (143) determined MIC and MBC values for amoxicillin, erythromycin, lincomycin, tetracycline, oxytetracycline, and minocycline for the infecting strain. Amoxicillin and erythromycin were the most effective, with MBCs of 0.5 µg/mL and 0.1 µg/mL, respectively. Recently, the susceptibilities of 11 serovars (seven species) of Leptospira to 14 antibiotics have been reported (144). With the exception of chloramphenicol, all tested agents were at least as potent as penicillin and doxycyline, with the macrolide and ketolide drugs producing the lowest MICs (≤0.01 µg/mL).

All of the organisms considered in this chapter, with the exception of the mycoplasmas, B. burgdorferi, and the Leptospira species, require either tissue culture or in vivo techniques for their propagation and susceptibility testing. Because of this, they frequently are not isolated from patients in whom they are causing disease.

If emerging resistance is to be detected, it is essential that recent clinical isolates are also tested, particularly those from patients who appear to be failing, or have failed, appropriate therapy. Our new real-time PCR assay could be useful in the future for the determination of the susceptibility to antibiotics of such fastidious bacteria (81). Moreover, recent development of new axenic culture media will also probably help to develop new, rapid, and simple antibiotic susceptibility assays in order to encourage laboratories to monitor the emergence of resistance for these intracellular bacteria in the future (8).

  1.  Senterfit LB. Antibiotic sensitivity testing of mycoplasmas. In: Razin S, Tully JG, eds. Methods in mycoplasmology. New York: Academic Press, 1983:397–401.

  2.  Berger BW, Kaplan MH, Rothenberg IR, et al. Isolation and characterization of the Lyme disease spirochete from the skin of patients with erythema chronicum migrans. J Am Acad Dermatol 1985;13(3):444–449.

  3.  Fieldsteel AH, Cox DL, Moeckli RA. Cultivation of virulent Treponema pallidum in tissue culture. Infect Immun 1981;32(2):908–915.

  4.  Stapleton JT, Stamm LV, Bassford PJ Jr. Potential for development of antibiotic resistance in pathogenic treponemes. Rev Infect Dis 1985;7(Suppl 2):S314–S317.

  5.  Stamm WE. Potential for antimicrobial resistance in Chlamydia pneumoniae. J Infect Dis 2000;181(Suppl 3):S456–S459.

  6.  Ehret JM, Judson FN. Susceptibility testing of Chlamydia trachomatis: from eggs to monoclonal antibodies. Antimicrob Agents Chemother 1988;32:1295–1299.

  7.  Ridgway GL, Bebear C, Bebear CM, et al. Antimicrobial susceptibility testing of intracellular and cell-associated pathogens. Clin Microbiol Infect 2001;7(12):1–10.

  8.  Singh S, Eldin C, Kowalczewska M, et al. Axenic culture of fastidious and intracellular bacteria. Trends Microbiol 2012;21(2):92–99.

  9.  McCormack WM. Susceptibility of mycoplasmas to antimicrobial agents: clinical implications. Clin Infect Dis 1993;17(Suppl 1):S200–S201.

 10.  Jones RB, Van der Pol B, Martin DH, et al. Partial characterization of Chlamydia trachomatis isolates resistant to multiple antibiotics. J Infect Dis 1990;162(6):1309–1315.

 11.  Mourad A, Sweet RL, Sugg N, et al. Relative resistance to erythromycin in Chlamydia trachomatis. Antimicrob Agents Chemother 1980;18(5):696–698.

 12.  Johnson FW, Clarkson MJ, Spencer WN. Susceptibility of Chlamydia psittaci (ovis) to antimicrobial agents. J Antimicrob Chemother 1983;11(5):413–418.

 13.  Grayston JT, Kuo CC, Wang SP, et al. A new Chlamydia psittaci strain, TWAR, isolated in acute respiratory tract infections. N Engl J Med 1986;315(3):161–168.

 14.  Thompson SE, Washington AE. Epidemiology of sexually transmitted Chlamydia trachomatis infections. Epidemiol Rev 1983;5:96–123.

 15.  Crosse BA. Psittacosis: a clinical review. J Infect 1990;21(3):251–259.

 16.  Kuo CC, Grayston JT. In vitro drug susceptibility of Chlamydia sp. strain TWAR. Antimicrob Agents Chemother 1988;32:257–258.

 17.  Cross NA, Kellock DJ, Kinghorn GR, et al. Antimicrobial susceptibility testing of Chlamydia trachomatis using a reverse transcriptase PCR-based method. Antimicrob Agents Chemother 1999;43(9):2311–2313.

 18.  Dessus-Babus S, Belloc F, Bebear CM, et al. Antibiotic susceptibility testing for Chlamydia trachomatis using flow cytometry. Cytometry 1998;31(1):37–44.

 19.  Bailey JM, Heppleston C, Richmond SJ. Comparison of the in vitro activities of ofloxacin and tetracycline against Chlamydia trachomatis as assessed by indirect immunofluorescence. Antimicrob Agents Chemother 1984;26(1):13–16.

 20.  Wyrick PB, Davis CH, Raulston JE, et al. Effect of clinically relevant culture conditions on antimicrobial susceptibility of Chlamydia trachomatis. Clin Infect Dis 1994;19(5):931–936.

 21.  Paul TR, Knight ST, Raulston JE, et al. Delivery of azithromycin to Chlamydia trachomatis-infected polarized human endometrial epithelial cells by polymorphonuclear leucocytes. J Antimicrob Chemother 1997;39(5):623–630.

 22.  Kutlin A, Roblin PM, Hammerschlag MR. In vitro activities of azithromycin and ofloxacin against Chlamydia pneumoniae in a continuous-infection model. Antimicrob Agents Chemother 1999;43(9):2268–2272.

 23.  Roblin PM, Kutlin A, Reznik T, et al. Activity of grepafloxacin and other fluoroquinones and newer macrolides against recent clinical isolates of Chlamydia pneumoniae. Int J Antimicrob Agents 1999;12(2):181–184.

 24.  Roblin PM, Reznik T, Kutlin A, et al. In vitro activities of gemifloxacin (SB 265805, LB20304) against recent clinical isolates of Chlamydia pneumoniae. Antimicrob Agents Chemother 1999;43(11):2806–2807.

 25.  Suchland RJ, Geisler WM, Stamm WE. Methodologies and cell lines used for antimicrobial susceptibility testing of Chlamydia spp. Antimicrob Agents Chemother 2003;47(2):636–642.

 26.  Hammerschlag MR. Activity of trimethoprim-sulfamethoxazole against Chlamydia trachomatis in vitro. Rev Infect Dis 1982;4(2):500–505.

 27.  Nagayama A, Nakao T, Taen H. In vitro activities of ofloxacin and four other new quinoline-carboxylic acids against Chlamydia trachomatis. Antimicrob Agents Chemother 1988;32(11):1735–1737.

 28.  Chirgwin K, Roblin PM, Hammerschlag MR. In vitro susceptibilities of Chlamydia pneumoniae (Chlamydia sp. strain TWAR). Antimicrob Agents Chemother 1989;33(9):1634–1635.

 29.  How SJ, Hobson D, Hart CA, et al. A comparison of the in-vitro activity of antimicrobials against Chlamydia trachomatis examined by Giemsa and a fluorescent antibody stain. J Antimicrob Chemother 1985;15(4):399–404.

 30.  Bowie WR. In vitro activity of clavulanic acid, amoxicillin, and ticarcillin against Chlamydia trachomatis. Antimicrob Agents Chemother 1986;29(4):713–715.

 31.  Chirgwin K, Roblin PM, Hammerschlag MR. In vitro susceptibilities of Chlamydia pneumoniae (Chlamydia sp. strain TWAR). J Antimicrob Chemother 1989;33:1634–1635.

 32.  Hammerschlag MR, Roblin PM, Bebear CM. Activity of telithromycin, a new ketolide antibacterial, against atypical and intracellular respiratory tract pathogens. J Antimicrob Chemother 2001;48(Suppl T1):25–31.

 33.  Roblin PM, Hammerschlag MR. In vitro activity of GAR-936 against Chlamydia pneumoniae and Chlamydia trachomatis. Int J Antimicrob Agents 2000;16(1):61–63.

 34.  Roblin PM, Hammerschlag MR. In vitro activity of a new antibiotic, NVP-PDF386 (VRC4887), against Chlamydia pneumoniae. Antimicrob Agents Chemother 2003;47(4):1447–1448.

 35.  Roblin PM, Reznik T, Kutlin A, et al. In vitro activities of rifamycin derivatives ABI-1648 (Rifalazil, KRM-1648), ABI-1657, and ABI-1131 against Chlamydia trachomatis and recent clinical isolates of Chlamydia pneumoniae. Antimicrob Agents Chemother2003;47(3):1135–1136.

 36.  Segreti J, Kapell KS. In vitro activity of dirithromycin against Chlamydia trachomatis. Antimicrob Agents Chemother 1994;38(9):2213–2214.

 37.  Segreti J, Gvazdinskas L, Trenholme G. In vitro activity of minocycline and rifampin against staphylococci. Diagn Microbiol Infect 1989;12:253–255.

 38.  Cevenini R, Donati M, Sambri V, et al. Enzyme-linked immunosorbent assay for the in-vitro detection of sensitivity of Chlamydia trachomatis to antimicrobial drugs. J Antimicrob Chemother 1987;20(5):677–684.

 39.  Bianchi A, Scieux C, Salmeron CM, et al. Rapid determination of MICs of 15 antichlamydial agents by using an enzyme immunoassay (Chlamydiazyme). Antimicrob Agents Chemother 1988;32(9):1350–1353.

 40.  Khan MA, Potter CW, Sharrard RM. A reverse transcriptase-PCR based assay for in-vitro antibiotic susceptibility testing of Chlamydia pneumoniae. J Antimicrob Chemother 1996;37(4):677–685.

 41.  Schachter J. Rifampin in chlamydial infections. Rev Infect Dis 1983;5(Suppl 3): S562–S564.

 42.  Keshishyan H, Hanna L, Jawetz E. Emergence of rifampin-resistance in Chlamydia trachomatis. Nature 1973;244(5412):173–174.

 43.  Misyurina OY, Chipitsyna EV, Finashutina YP, et al. Mutations in a 23S rRNA gene of Chlamydia trachomatis associated with resistance to macrolides. Antimicrob Agents Chemother 2004;48(4):1347–1349.

 44.  Lenart J, Andersen AA, Rockey DD. Growth and development of tetracycline-resistant Chlamydia suis. Antimicrob Agents Chemother 2001;45(8):2198–2203.

 45.  Sandoz KM, Rockey DD. Antibiotic resistance in Chlamydiae. Future Microbiol 2010;5(9):1427–1442.

 46.  Dessus-Babus S, Bebear CM, Charron A, et al. Sequencing of gyrase and topoisomerase IV quinolone-resistance-determining regions of Chlamydia trachomatis and characterization of quinolone-resistant mutants obtained In vitro. Antimicrob Agents Chemother1998;42(10):2474–2481.

 47.  Lo SC, Wear DJ, Green SL, et al. Adult respiratory distress syndrome with or without systemic disease associated with infections due to Mycoplasma fermentans. Clin Infect Dis 1993;17(Suppl 1):S259–S263.

 48.  Senterfit LB. Laboratory diagnosis of mycoplasma infections. Isr J Med Sci 1984;20(10):905–907.

 49.  Kenny GE, Cartwright FD, Roberts MC. Agar dilution method for determination of antibiotic susceptibility of Ureaplasma urealyticum. Pediatr Infect Dis 1986;5(Suppl 6):S332–S334.

 50.  Waites KB, Crabb DM, Bing X, et al. In vitro susceptibilities to and bactericidal activities of garenoxacin (BMS-284756) and other antimicrobial agents against human mycoplasmas and ureaplasmas. Antimicrob Agents Chemother 2003;47(1):161–165.

 51.  Waites KB, Figarola TA, Schmid T, et al. Comparison of agar versus broth dilution techniques for determining antibiotic susceptibilities of Ureaplasma urealyticum. Diagn Microbiol Infect Dis 1991;14(3):265–271.

 52.  Roberts MC, Koutsky LA, Holmes KK, et al. Tetracycline-resistant Mycoplasma hominis strains contain streptococcal tetM sequences. Antimicrob Agents Chemother 1985;28(1):141–143.

 53.  Tanner AC, Erickson BZ, Ross RF. Adaptation of the Sensititre broth microdilution technique to antimicrobial susceptibility testing of Mycoplasma hyopneumoniae. Vet Microbiol 1993;36(3–4):301–306.

 54.  Poulin SA, Perkins RE, Kundsin RB. Antibiotic susceptibilities of AIDS-associated mycoplasmas. J Clin Microbiol 1994;32(4):1101–1103.

 55.  Limb DI, Wheat PF, Hastings JG, et al. Antimicrobial susceptibility testing of mycoplasmas by ATP bioluminescence. J Med Microbiol 1991;35(2):89–92.

 56.  Tully JG, Whitcomb RF, Clark HF, et al. Pathogenic mycoplasmas: cultivation and vertebrate pathogenicity of a new spiroplasma. Science 1977;195(4281):892–894.

 57.  Waites KB, Cassell GH, Canupp KC, et al. In vitro susceptibilities of mycoplasmas and ureaplasmas to new macrolides and aryl-fluoroquinolones. Antimicrob Agents Chemother 1988;32(10):1500–1502.

 58.  Waites KB, Crouse DT, Nelson KG, et al. Chronic Ureaplasma urealyticum and Mycoplasma hominis infections of central nervous system in preterm infants. Lancet 1988;1(8575–8576):17–21.

 59.  Robertson JA, Coppola JE, Heisler OR. Standardized method for determining antimicrobial susceptibility of strains of Ureaplasma urealyticum and their response to tetracycline, erythromycin, and rosaramicin. Antimicrob Agents Chemother 1981;20(1):53–58.

 60.  Taylor-Robinson D, Furr PM. The static effect of rosaramicin on Ureaplasma urealyticum and the development of antibiotic resistance. J Antimicrob Chemother 1982;10(3):185–191.

 61.  Rylander M, Hallander HO. In vitro comparison of the activity of doxycycline, tetracycline, erythromycin and a new macrolide, CP 62993, against Mycoplasma pneumoniae, Mycoplasma hominis and Ureaplasma urealyticum. Scand J Infect Dis Suppl 1988;53:12–17.

 62.  Pereyre S, Guyot C, Renaudin H, et al. In vitro selection and characterization of resistance to macrolides and related antibiotics in Mycoplasma pneumoniae. Antimicrob Agents Chemother 2004;48(2):460–465.

 63.  Stopler T, Branski D. Resistance of Mycoplasma pneumoniae to macrolides, lincomycin and streptogramin B. J Antimicrob Chemother 1986;18(3):359–364.

 64.  Bebear C, Pereyre S, Peuchant O. Mycoplasma pneumoniae: susceptibility and resistance to antibiotics. Future Microbiol 2011;6(4):423–431.

 65.  Ishida K, Kaku M, Irifune K, et al. In vitro and in vivo activities of macrolides against Mycoplasma pneumoniae. Antimicrob Agents Chemother 1994;38(4):790–798.

 66.  Yamaguchi T, Hirakata Y, Izumikawa K, et al. In vitro activity of telithromycin (HMR3647), a new ketolide, against clinical isolates of Mycoplasma pneumoniae in Japan. Antimicrob Agents Chemother 2000;44(5):1381–1382.

 67.  Hammerschlag MR, Hyman CL, Roblin PM. In vitro activities of five quinolones against Chlamydia pneumoniae. Antimicrob Agents Chemother 1992;36:682–683.

 68.  Kaku M, Ishida K, Irifune K, et al. In vitro and in vivo activities of sparfloxacin against Mycoplasma pneumoniae. Antimicrob Agents Chemother 1994;38(4):738–741.

 69.  Kenny GE, Cartwright FD. Susceptibility of Mycoplasma pneumoniae to several new quinolones, tetracycline, and erythromycin. Antimicrob Agents Chemother 1991;35(3):587–589.

 70.  Krausse R, Schubert S. In-vitro activities of tetracyclines, macrolides, fluoroquinolones and clindamycin against Mycoplasma hominis and Ureaplasma ssp. isolated in Germany over 20 years. Clin Microbiol Infect 2010;16(11):1649–1655.

 71.  Taylor-Robinson D, Furr PM. Clinical antibiotic resistance of Ureaplasma urealyticum. Pediatr Infect Dis 1986;5(6)(Suppl):S335–S337.

 72.  Waites KB, Crouse DT, Cassell GH. Antibiotic susceptibilities and therapeutic options for Ureaplasma urealyticum infections in neonates. Pediatr Infect Dis J 1992;11(1):23–29.

 73.  Waites KB, Crouse DT, Cassell GH. Therapeutic considerations for Ureaplasma urealyticum infections in neonates. Clin Infect Dis 1993;17(Suppl 1):S208–S214.

 74.  Bebear CM, Bove JM, Bebear C, et al. Characterization of Mycoplasma hominis mutations involved in resistance to fluoroquinolones. Antimicrob Agents Chemother 1997;41(2):269–273.

 75.  MacKenzie CR, Nischik N, Kram R, et al. Fatal outcome of a disseminated dual infection with drug-resistant Mycoplasma hominis and Ureaplasma parvum originating from a septic arthritis in an immunocompromised patient. Int J Infect Dis 2010;14(Suppl 3):e307–e309.

 76.  Pereyre S, Gonzalez P, de Barbeyrac B, et al. Mutations in 23S rRNA account for intrinsic resistance to macrolides in Mycoplasma hominis and Mycoplasma fermentans and for acquired resistance to macrolides in M. hominis. Antimicrob Agents Chemother 2002;46(10):3142–3150.

 77.  Parola P, Paddock CD, Raoult D. Tick-borne rickettsioses around the world: emerging diseases challenging old concepts. Clin Microbiol Rev 2005;18(4):719–756.

 78.  Raoult D, Roussellier P, Vestris G, et al. In vitro antibiotic susceptibility of Rickettsia rickettsii and Rickettsia conorii: plaque assay and microplaque colorimetric assay. J Infect Dis 1987;155:1059–1062.

 79.  Raoult D, Roussellier P, Galicher V, et al. In vitro susceptibility of Rickettsia conorii to ciprofloxacin as determined by suppressing lethality in chicken embryos and by plaque assay. Antimicrob Agents Chemother 1986;29:424–425.

 80.  Rolain JM, Maurin M, Vestris G, et al. In vitro susceptibilities of 27 Rickettsiae to 13 antimicrobials. Antimicrob Agents Chemother 1998;42(7):1537–1541.

 81.  Rolain JM, Stuhl L, Maurin M, et al. Evaluation of antibiotic susceptibilities of three rickettsial species including Rickettsia felis by a quantitative PCR DNA assay. Antimicrob Agents Chemother 2002;46(9):2747–2751.

 82.  Ives TJ, Marston EL, Regnery RL, et al. In vitro susceptibilities of Bartonella and Rickettsia spp. to fluoroquinolone antibiotics as determined by immunofluorescent antibody analysis of infected Vero cell monolayers. Int J Antimicrob Agents 2001;18(3):217–222.

 83.  Ives TJ, Manzewitsch P, Regnery RL, et al. In vitro susceptibilities of Bartonella henselae, B. quintana, B. elizabethae, Rickettsia rickettsii, R. conorii, R. akari, and R. prowazekii to macrolide antibiotics as determined by immunofluorescent-antibody analysis of infected vero cell monolayers. Antimicrob Agents Chemother 1997;41:578–582.

 84.  Raoult D, Roussellier P, Tamalet J. In vitro evaluation of josamycin, spiramycin, and erythromycin against Rickettsia rickettsii and R. conorii. Antimicrob Agents Chemother 1988;32:255–256.

 85.  McLaren C, Ellis MN, Hunter GA. A colorimetric assay for the measurement of the sensitivity of herpes simplex viruses to antiviral agents. Antiviral Res 1983;3(4):223–234.

 86.  Wittwer CT, Ririe KM, Andrew RV, et al. The LightCycler: a microvolume multisample fluorimeter with rapid temperature control. Biotechniques 1997;22(1):176–181.

 87.  Rolain JM, Raoult D. Prediction of resistance to erythromycin in the genus Rickettsia by mutations in L22 ribosomal protein. J Antimicrob Chemother 2005;56(2):396–398.

 88.  Drancourt M, Raoult D. Characterization of mutations in the rpoB gene in naturally rifampin-resistant Rickettsia species. Antimicrob Agent Chemother 1999;43(10):2400–2403.

 89.  Dumler JS, Barbet AF, Bekker CP, et al. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and ‘HGE agent’ as subjective synonyms of Ehrlichia phagocytophila. Int J Syst Evol Microbiol 2001;51(Pt 6):2145–2165.

 90.  Dumler JS, Bakken JS. Ehrlichial diseases of humans: emerging tick-borne infections. Clin Infect Dis 1995;20:1102–1110.

 91.  Tachibana N, Kobayashi V. Effect of cyclophosphamide on the growth of Rickettsia sennetsu in experimentally infected mice. Infect Immun 1975;12:625–629.

 92.  Kobayashi Y, Ikeda O, Miaso T. Chemotherapy of sennetsu disease. Progress in virology. Tokyo, Japan: Bainukan, 1962:130–142.

 93.  Kelly DJ, LaBarre DD, Lewis GEJ. Effect of tetracycline therapy on host defense in mice infected with Ehrlichia sennetsu. In: Winkler HH, Ristic M, eds. Microbiology. Washington, DC: American Society for Microbiology, 1986:209–212.

 94.  Oyama T. Immunological studies of rickettsial infection—analysis of lymphoid cell subpopulations of the spleen of mice infected with Rickettsia sennetsu and Rickettsia tsutsugamushi [in Japanese]. Kansenshogaku Zasshi 1979;53:243–257.

 95.  Brouqui P, Raoult D. In vitro susceptibility of Ehrlichia sennetsu to antibiotics. Antimicrob Agents Chemother 1990;34:1593–1596.

 96.  Brouqui P, Raoult D. In vitro antibiotic susceptibility of the newly recognized agent of ehrhlichiosis in humans, Ehrlichia chaffeensis. Antimicrob Agents Chemother 1992;36:2799–2803.

 97.  Rolain JM, Maurin M, Bryskier A, et al. In vitro activities of telithromycin (HMR 3647) against Rickettsia rickettsii, Rickettsia conorii, Rickettsia africae, Rickettsia typhi, Rickettsia prowasekii, Coxiella burnetii, Bartonella henselae, Bartonella quintana, Bartonella bacilliformis, and Ehrlichia chaffeensis. Antimicrob Agents Chemother 2000;44(5):1391–1393.

 98.  Horowitz HW, Hsieh TC, Aguero-Rosenfeld ME, et al. Antimicrobial susceptibility of Ehrlichia phagocytophila. Antimicrob Agents Chemother 2001;45(3):786–788.

 99.  Klein MB, Nelson CM, Goodman JL. Antibiotic susceptibility of the newly cultivated agent of human granulocytic ehrlichiosis: promising activity of quinolones and rifamycins. Antimicrob Agents Chemother 1997;41:76–79.

100.  Maurin M, Bakken JS, Dumler JS. Antibiotic susceptibilities of Anaplasma (Ehrlichia) phagocytophilum strains from various geographic areas in the United States. Antimicrob Agents Chemother 2003;47(1):413–415.

101.  Maurin M, Abergel C, Raoult D. DNA Gyrase-mediated natural resistance to fluoroquinolones in Ehrlichia spp. Antimicrob Agents Chemother 2001;45(7):2098–2105.

102.  Branger S, Rolain JM, Raoult D. Evaluation of antibiotic susceptibilities of Ehrlichia canis, Ehrlichia chaffeensis, and Anaplasma phagocytophilum by real-time PCR. Antimicrob Agents Chemother 2004;48(12):4822–4828.

103.  Tantibhedhyangkul W, Angelakis E, Tongyoo N, et al. Intrinsic fluoroquinolone resistance in Orientia tsutsugamushi. Int J Antimicrob Agents 2010;35(4):338–341.

104.  Maurin M, Benoliel AM, Bongrand P, et al. Phagolysosomes of Coxiella burnetii-infected cell lines maintain an acidic pH during persistent infection. Infect Immun 1992;60(12):5013–5016.

105.  Maurin M, Raoult D. Q fever. Clin Microbiol Rev 1999;12(4):518–553.

106.  Christopher GW, Cieslak TJ, Pavlin JA, et al. Biological warfare. A historical perspective. JAMA 1997;278(5):412–417.

107.  Greenfield RA, Drevets DA, Machado LJ, et al. Bacterial pathogens as biological weapons and agents of bioterrorism. Am J Med Sci 2002;323(6):299–315.

108.  Raoult D, Mege JL, Marrie T. Q fever: queries remaining after decades of research. In: Scheld WM, Craig WA, Hughes JM, eds. Emerging infections. Washington, DC: ASM Press, 2001:29–56.

109.  Boulos A, Rolain JM, Maurin M, et al. Evaluation of antibiotic susceptibilities against Coxiella burnetii by real time PCR. Int J Antimicrob Agents 2004;23(2):169–174.

110.  Brennan RE, Samuel JE. Evaluation of Coxiella burnetii antibiotic susceptibilities by real-time PCR assay. J Clin Microbiol 2003;41(5):1869–1874.

111.  Rolain JM, Lambert F, Raoult D. Activity of telithromycin against thirteen new isolates of C. burnetii including three resistant to doxycycline. Ann N Y Acad Sci 2006;1063:252–256.

112.  Beare PA, Sandoz KM, Omsland A, et al. Advances in genetic manipulation of obligate intracellular bacterial pathogens. Front Microbiol 2011;2:97.

113.  Omsland A, Cockrell DC, Howe D, et al. Host cell-free growth of the Q fever bacterium Coxiella burnetii. Proc Natl Acad Sci U S A 2009;106(11):4430–4434.

114.  Jabarit-Aldighieri N, Torres H, Raoult D. Susceptibility of R. conorii, R.rickettsii and C.burnetii to CI- 960 (PD 127,391), PD 131,628, pefloxacin, ofloxacin and ciprofloxacin. Antimicrob Agents Chemother 1992;36:2529–2532.

115.  Maurin M, Raoult D. In vitro susceptibilities of spotted fever group rickettsiae and Coxiella burnetii to clarithromycin. Antimicrob Agents Chemother 1993;37:2633–2637.

116.  Spyridaki I, Psaroulaki A, Vranakis I, et al. Bacteriostatic and bactericidal activities of tigecycline against Coxiella burnetii and comparison with those of six other antibiotics. Antimicrob Agents Chemother 2009;53(6):2690–2692.

117.  Raoult D, Torres H, Drancourt M. Shell-vial assay: evaluation of a new technique for determining antibiotic susceptibility, tested in 13 isolates of Coxiella burnetii. Antimicrob Agents Chemother 1991;35(10):2070–2077.

118.  Torres H, Raoult D. In vitro activities of ceftriaxone and fusidic acid against 13 isolates of Coxiella burnetii, determined using the shell vial assay. Antimicrob Agents Chemother 1993;37:491–494.

119.  Baca OG, Akporiaye ET, Aragon AS, et al. Fate of phase I and phase II Coxiella burnetii in several macrophage-like tumor cell lines. Infect Immun 1981;33:258–266.

120.  Roman MJ, Coriz PD, Baca OG. A proposed model to explain persistent infection of host cells with Coxiella burnetii. J Gen Microbiol 1986;132:1415–1422.

121.  Raoult D, Drancourt M, Vestris G. Bactericidal effect of doxycycline associated with lysosomotropic agents on Coxiella burnetii in P388D1 cells. Antimicrob Agents Chemother 1990;34:1512–1514.

122.  Rouli L, Rolain JM, El Filali A, et al. Genome sequence of Coxiella burnetii 109, a doxycycline-resistant clinical isolate. J Bacteriol 2012;194(24):6939.

123.  Maurin M, Benoliel AM, Bongrand P, et al. Phagolysosomal alkalinization and the bactericidal effect of antibiotics: the Coxiella burnetii paradigm. J Infect Dis 1992;166:1097–1102.

124.  Raoult D, Vestris G, Enea M. Isolation of 16 strains of Coxiella burnetii from patients by using a sensitive centrifugation cell culture system and establishment of strains in HEL cells. J Clin Microbiol 1990;28:2482–2484.

125.  Agger WA, Callister SM, Jobe DA. In vitro susceptibilities of Borrelia burgdorferi to five oral cephalosporins and ceftriaxone. Antimicrob Agents Chemother 1992;36(8):1788–1790.

126.  Norris SJ, Edmondson DG. In vitro culture system to determine MICs and MBCs of antimicrobial agents against Treponema pallidum subsp. pallidum (Nichols strain). Antimicrob Agents Chemother 1988;32(1):68–74.

127.  Dever LL, Jorgensen JH, Barbour AG. In vitro antimicrobial susceptibility testing of Borrelia burgdorferi: a microdilution MIC method and time-kill studies. J Clin Microbiol 1992;30(10):2692–2697.

128.  Preac-Mursic V, Wilske B, Schierz G. European Borrelia burgdorferi isolated from humans and ticks culture conditions and antibiotic susceptibility. Zentralbl Bakteriol Mikrobiol Hyg A 1986;263(1–2):112–118.

129.  Donati M, Pollini GM, Sparacino M, et al. Comparative in vitro activity of garenoxacin against Chlamydia spp. J Antimicrob Chemother 2002;50(3):407–410.

130.  Hunfeld KP, Kraiczy P, Wichelhaus TA, et al. Colorimetric in vitro susceptibility testing of penicillins, cephalosporins, macrolides, streptogramins, tetracyclines, and aminoglycosides against Borrelia burgdorferi isolates. Int J Antimicrob Agents 2000;15(1):11–17.

131.  Sapi E, Kaur N, Anyanwu S, et al. Evaluation of in-vitro antibiotic susceptibility of different morphological forms of Borrelia burgdorferi. Infect Drug Resist 2011;4:97–113.

132.  Levin JM, Nelson JA, Segreti J, et al. In vitro susceptibility of Borrelia burgdorferi to 11 antimicrobial agents. Antimicrob Agents Chemother 1993;37(7):1444–1446.

133.  Johnson SE, Klein GC, Schmid GP, et al. Susceptibility of the Lyme disease spirochete to seven antimicrobial agents. Yale J Biol Med 1984;57(4):549–553.

134.  Levy JA. Confirmation of the successful cultivation of Treponema pallidum in tissue culture. Microbiologica 1984;7:367–370.

135.  Stamm LV, Stapleton JT, Bassford PJ Jr. In vitro assay to demonstrate high-level erythromycin resistance of a clinical isolate of Treponema pallidum. Antimicrob Agents Chemother 1988;32(2):164–169.

136.  Stamm LV. Global challenge of antibiotic-resistant Treponema pallidum. Antimicrob Agents Chemother 2010;54(2):583–589.

137.  Johnson RC, Harris VG. Differentiation of pathogenic and saprophytic letospires. I. Growth at low temperatures. J Bacteriol 1967;94(1):27–31.

138.  Stamm LV, Bergen HL. A point mutation associated with bacterial macrolide resistance is present in both 23S rRNA genes of an erythromycin-resistant Treponema pallidum clinical isolate. Antimicrob Agents Chemother 2000;44(3):806–807.

139.  Levett PN. Leptospirosis. Clin Microbiol Rev 2001;14(2):296–326.

140.  Arzouni JP, Parola P, La Scola B, et al. Human infection caused by Leptospira fainei. Emerg Infect Dis 2002;8(8):865–868.

141.  Petersen AM, Boye K, Blom J, et al. First isolation of Leptospira fainei serovar Hurstbridge from two human patients with Weil’s syndrome. J Med Microbiol 2001;50(1):96–100.

142.  Oie S, Hironaga K, Koshiro A, et al. In vitro susceptibilities of five Leptospira strains to 16 antimicrobial agents. Antimicrob Agents Chemother 1983;24(6):905–908.

143.  Broughton ES, Flack LE. The susceptibility of a strain of Leptospira interrogans serogroup icterohaemorrhagiae to amoxycillin, erythromycin, lincomycin, tetracycline, oxytetracycline and minocycline. Zentralbl Bakteriol Mikrobiol Hyg A 1986;261(4):425–431.

144.  Hospenthal DR, Murray CK. In vitro susceptibilities of seven Leptospira species to traditional and newer antibiotics. Antimicrob Agents Chemother 2003;47(8):2646–2648.

145.  Bowie WR, Lee CK, Alexander ER. Prediction of efficacy of antimicrobial agents in treatment of infections due to Chlamydia trachomatis. J Infect Dis 1978;138(5):655–659.

146.  Johannisson G, Sernryd A, Lycke E. Susceptibility of Chlamydia trachomatis to antibiotics in vitro and in vivo. Sex Transm Dis 1979;6(2):50–57.

147.  Zurenko GE, Yagi BH, Vavra JJ, et al. In vitro antibacterial activity of trospectomycin (U-63366F), a novel spectinomycin analog. Antimicrob Agents Chemother 1988;32(2):216–223.

148.  Bowie WR. Lack of in vitro activity of cefoxitin, cefamandole, cefuroxime, and piperacillin against Chlamydia trachomatis. Antimicrob Agents Chemother 1982;21(2):339–340.

149.  Hammerschlag MR, Gleyzer A. In vitro activity of a group of broad spectrum cephalosporins and other betalactam antibodies against Chlamydia trachomatis. Antimicrob Agents Chemother 1983;23:493–494.

150.  Martin DH, Pastorek JG, Faro S. In-vitro and in-vivo activity of parenterally administered beta-lactam antibiotics against Chlamydia trachomatis. Sex Transm Dis 1986;13(2):81–87.

151.  Muytjens HL, Heessen FW. In vitro activities of thirteen beta-lactam antibiotics against Chlamydia trachomatis. Antimicrob Agents Chemother 1982;22(3):520–521.

152.  Agacfidan A, Moncada J, Schachter J. In vitro activity of azithromycin (CP-62,993) against Chlamydia trachomatis and Chlamydia pneumoniae. Antimicrob Agents Chemother 1993;37(9):1746–1748.

153.  Rumpianesi F, Morandotti G, Sperning R, et al. In vitro activity of azithromycin against Chlamydia trachomatis, Ureaplasma urealyticum and Mycoplasma hominis in comparison with erythromycin, roxithromycin and minocycline. J Chemother 1993;5(3):155–158.

154.  Welsh LE, Gaydos CA, Quinn TC. In vitro evaluation of activities of azithromycin, erythromycin, and tetracycline against Chlamydia trachomatis and Chlamydia pneumoniae. Antimicrob Agents Chemother 1992;36:291–294.

155.  Benson C, Segreti J, Kessler H, et al. Comparative in vitro activity of A-56268 (TE-031) against gram-positive and gram-negative bacteria and Chlamydia trachomatis. Eur J Clin Microbiol 1987;6(2):173–178.

156.  Ridgway GL, Mumtaz G, Fenelon L. The in-vitro activity of clarithromycin and other macrolides against the type strain of Chlamydia pneumoniae (TWAR). J Antimicrob Chemother 1991;27(Suppl A):43–45.

157.  Samra Z, Rosenberg S, Soffer Y, et al. In vitro susceptibility of recent clinical isolates of Chlamydia trachomatis to macrolides and tetracyclines. Diagn Microbiol Infect Dis 2001;39(3):177–179.

158.  Stamm WE, Suchland R. Antimicrobial activity of U-70138F (paldimycin), roxithromycin (RU 965), and ofloxacin (ORF 18489) against Chlamydia trachomatis in cell culture. Antimicrob Agents Chemother 1986;30(5):806–807.

159.  Miyashita N, Fukano H, Niki Y, et al. In vitro activity of telithromycin, a new ketolide, against Chlamydia pneumoniae. J Antimicrob Chemother 2001;48(3):403–405.

160.  Miyashita N, Fukano H, Yoshida K, et al. In vitro activity of cethromycin, a novel antibacterial ketolide, against Chlamydia pneumoniae. J Antimicrob Chemother 2003;52(3):497–499.

161.  Kuo CC, Wang SP, Grayston JT. Antimicrobial activity of several antibiotics and a sulfonamide against Chlamydia trachomatis organisms in cell culture. Antimicrob Agents Chemother 1977;12(1):80–83.

162.  Gieffers J, Solbach W, Maass M. In vitro susceptibilities of Chlamydia pneumoniae strains recovered from atherosclerotic coronary arteries. Antimicrob Agents Chemother 1998;42(10):2762–2764.

163.  Miyashita N, Fukano H, Yoshida K, et al. In-vitro activity of moxifloxacin and other fluoroquinolones against Chlamydia species. J Infect Chemother 2002;8(1):115–117.

164.  Maeda H, Fujii A, Nakata K, et al. In vitro activities of T-3262, NY-198, fleroxacin (AM-833; RO 23-6240), and other new quinolone agents against clinically isolated Chlamydia trachomatis strains. Antimicrob Agents Chemother 1988;32(7):1080–1081.

165.  Roblin PM, Reznik T, Hammerschlag MR. In vitro activity of garenoxacin against recent clinical isolates of Chlamydia pneumoniae. Int J Antimicrob Agents 2003;21(6):578–580.

166.  Roblin PM, Hammerschlag MR. In-vitro activity of gatifloxacin against Chlamydia trachomatis and Chlamydia pneumoniae. J Antimicrob Chemother 1999;44(4):549–551.

167.  Hammerschlag MR. Activity of gemifloxacin and other new quinolones against Chlamydia pneumoniae: a review. J Antimicrob Chemother 2000;45(Suppl 1):35–39.

168.  Heessen FW, Muytjens HL. In vitro activities of ciprofloxacin, norfloxacin, pipemidic acid, cinoxacin, and nalidixic acid against Chlamydia trachomatis. Antimicrob Agents Chemother 1984;25(1):123–124.

169.  Segreti J, Kessler HA, Kapell KS, et al. In vitro activities of temafloxacin (A-62254) and four other antibiotics against Chlamydia trachomatis. Antimicrob Agents Chemother 1989;33(1):118–119.

170.  Hardy DJ. Activity of temafloxacin and other fluoroquinolones against typical and atypical community-acquired respiratory tract pathogens. Am J Med 1991;91(6A):12S–14S.

171.  Lefevre JC, Bauriaud R, Gaubert E, et al. In vitro activity of sparfloxacin and other antimicrobial agents against genital pathogens. Chemotherapy 1992;38(5):303–307.

172.  Roblin PM, Kutlin A, Hammerschlag MR. In vitro activity of trovafloxacin against Chlamydia pneumoniae. Antimicrob Agents Chemother 1997;41(9):2033–2034.

173.  Jones RB, Van der Pol B, Johnson RB. Susceptibility of Chlamydia trachomatis to trovafloxacin. J Antimicrob Chemother 1997;39(Suppl B):63–65.

174.  Christensen JJ, Holten-Andersen W, Nielsen PB. Chlamydia trachomatis: in vitro susceptibility to antibiotics singly and in combination. Acta Pathol Microbiol Immunol Scand B 1986;94(5):329–332.

175.  Gnarpe J, Eriksson K, Gnarpe H. In vitro activities of azithromycin and doxycycline against 15 isolates of Chlamydia pneumoniae. Antimicrob Agents Chemother 1996;40(8):1843–1845.

176.  Kimura M, Kishimoto T, Niki Y, et al. In vitro and in vivo antichlamydial activities of newly developed quinolone antimicrobial agents. Antimicrob Agents Chemother 1993;37(4):801–803.

177.  Harrison HR, Riggin RM, Alexander ER, et al. In vitro activity of clindamycin against strains of Chlamydia trachomatis, Mycoplasma hominis, and Ureaplasma urealyticum isolated from pregnant women. Am J Obstet Gynecol 1984;149(5):477–480.

178.  Jones RB, Ridgway GL, Boulding S, et al. In vitro activity of rifamycins alone and in combination with other antibiotics against Chlamydia trachomatis. Rev Infect Dis 1983;5(Suppl 3):S556–S561.

179.  Freidank HM, Losch P, Vogele H, et al. In vitro susceptibilities of Chlamydia pneumoniae isolates from German patients and synergistic activity of antibiotic combinations. Antimicrob Agents Chemother 1999;43(7):1808–1810.

180.  Jenkin HM, Hung SC. Effect of vancomycin on the growth of psittacosis-trachoma agents cultivated in eggs and cell culture. Appl Microbiol 1967;15(1):10–12.

181.  Jao RL. Susceptibility of Mycoplasma pneumoniae to 21 antibiotics in vitro. Am J Med Sci 1967;253(6):639–650.

182.  Kenny GE, Cartwright FD. Susceptibilities of Mycoplasma hominis, M. pneumoniae, and Ureaplasma urealyticum to GAR-936, dalfopristin, dirithromycin, evernimicin, gatifloxacin, linezolid, moxifloxacin, quinupristin-dalfopristin, and telithromycin compared to their susceptibilities to reference macrolides, tetracyclines, and quinolones. Antimicrob Agents Chemother 2001;45(9):2604–2608.

183.  Renaudin H, Bebear C. Comparative in vitro activity of azithromycin, clarithromycin, erythromycin and lomefloxacin against Mycoplasma pneumoniae, Mycoplasma hominis and Ureaplasma urealyticum. Eur J Clin Microbiol Infect Dis 1990;9(11):838–841.

184.  Misu T, Arai S, Furukawa M, et al. Effects of rokitamycin and other macrolide antibiotics on Mycoplasma pneumoniae in L cells. Antimicrob Agents Chemother 1987;31:1843–1845.

185.  Yu X, Fan M, Xu G, et al. Genotypic and antigenic identification of two new strains of spotted fever group rickettsiae isolated from China. J Clin Microbiol 1993;31:83–88.

186.  Cassell GH, Waites KB, Pate MS, et al. Comparative susceptibility of Mycoplasma pneumoniae to erythromycin, ciprofloxacin, and lomefloxacin. Diagn Microbiol Infect Dis 1989;12(5):433–435.

187.  Kenny GE, Hooton TM, Roberts MC, et al. Susceptibilities of genital mycoplasmas to the newer quinolones as determined by the agar dilution method. Antimicrob Agents Chemother 1989;33(1):103–107.

188.  Osada Y, Ogawa H. Antimycoplasmal activity of ofloxacin (DL-8280). Antimicrob Agents Chemother 1983;23(3):509–511.

189.  Waites KB, Crabb DM, Duffy LB. Inhibitory and bactericidal activities of gemifloxacin and other antimicrobials against Mycoplasma pneumoniae. Int J Antimicrob Agents 2003;21(6):574–577.

190.  Kenny GE, Cartwright FD. Susceptibilities of Mycoplasma pneumoniae, Mycoplasma hominis, and Ureaplasma urealyticum to a new quinolone, trovafloxacin (CP-99,219). Antimicrob Agents Chemother 1996;40(4):1048–1049.

191.  Bygdeman SM, Mardh PA. Antimicrobial susceptibility and susceptibility testing of Mycoplasma hominis: a review. Sex Transm Dis 1983;10(4)(Suppl):366–370.

192.  Kenny GE, Cartwright FD. Susceptibilities of Mycoplasma hominis, Mycoplasma pneumoniae, and Ureaplasma urealyticum to new glycylcyclines in comparison with those to older tetracyclines. Antimicrob Agents Chemother 1994;38(11):2628–2632.

193.  Brouqui P, Raoult D. Susceptibilities of Ehrlichia to antibiotics. In: Raoult D, ed. Antimicrobial agents and intracellular pathogens. 9th ed. Boca Raton, FL: CRC Press, 1993:179–199.

194.  Ates L, Hanssen-Hubner C, Norris DE, et al. Comparison of in vitro activities of tigecycline, doxycycline, and tetracycline against the spirochete Borrelia burgdorferi. Ticks Tick Borne Dis 2010;1(1):30–34.

195.  Mursic VP, Wilske B, Schierz G, et al. In vitro and in vivo susceptibility of Borrelia burgdorferi. Eur J Clin Microbiol 1987;6(4):424–426.

196.  Johnson RC, Kodner C, Russell M. In vitro and in vivo susceptibility of the Lyme disease spirochete, Borrelia burgdorferi, to four antimicrobial agents. Antimicrob Agents Chemother 1987;31(2):164–167.

197.  Sicklinger M, Wienecke R, Neubert U. In vitro susceptibility testing of four antibiotics against Borrelia burgdorferi: a comparison of results for the three genospecies Borrelia afzelii, Borrelia garinii, and Borrelia burgdorferi sensu stricto. J Clin Microbiol 2003;41(4):1791–1793.

198.  Cinco M, Padovan D, Stinco G, et al. In vitro activity of rokitamycin, a new macrolide, against Borrelia burgdorferi. Antimicrob Agents Chemother 1995;39(5):1185–1186.

199.  Goldmeier D, Hay P. A review and update on adult syphilis, with particular reference to its treatment. Int J STD AIDS 1993;4(2):70–82.

200.  Korting HC, Walther D, Riethmuller U, et al. Comparative in vitro susceptibility of Treponema pallidum to ceftizoxime, ceftriaxone and penicillin G. Chemotherapy 1986;32(4):352–355.