Evaluation of Potential Chemotherapeutic Antimicrobials

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Chapter: Pharmaceutical Microbiology : Laboratory Evaluation Of Antimicrobial Agents

Unlike tests for the evaluation of disinfectants, where determination of cidal activity is of paramount importance, tests involving potential chemotherapeutic agents (antibiotics) invariably have determination of MIC as their main focus.



Unlike tests for the evaluation of disinfectants, where determination of cidal activity is of paramount importance, tests involving potential chemotherapeutic agents (antibiotics) invariably have determination of MIC as their main focus. Tests for the bacteriostatic activity of antimicrobial agents are valuable tools in predicting antimicrobial sensitivity/tolerance in individual patient samples and for detection and monitoring of resistant bacteria. However, correlation between MIC and therapeutic outcome are frequently difficult to predict, especially in chronic biofilm-mediated infections. The determination of MIC values must be conducted under standardized conditions, since deviation from standard test conditions can result in considerable variation in data.


a)     Tests For Bacteriostatic Activity


The historical gradient plates, ditch-plate and cup-plate techniques have been replaced by more quantitative techniques such as disc diffusion (Figure 18.4), broth and agar dilution, and E-tests (Figure 18.5). All employ chemically defined media (e.g. Mueller-Hinton or Iso-Sensitest) at a pH of 7.2–7.4, and in the case of solid media, agar plates of defined thickness. Regularly updated guidelines have been provided by the National Committee for Clinical Laboratory Standards (NCCLS) and are widely used in many countries, although the British Society for Antimicrobial Chemotherapy has produced its own guidelines and testing procedures (Andrews, 2009).



                i)  Disc tests


These are really modifications of the earlier cup or ditch plate procedures where filter-paper discs impregnated with the antimicrobial replace the antimicrobial-filled cups or wells. For disc tests, standard suspensions (e.g. 0.5 McFarland standard) of log-phase growth cells are prepared and inoculated on to the surface of appropriate agar plates to form a lawn. Commercially available filter paper discs containing known concentrations of antimicrobial agent (it is possible to prepare your own discs for use with novel drugs) are then placed on the dried lawn and the plates are incubated aerobically at 35 °C for 18 hours. The density of bacteria inoculated on to the plate should produce just confluent growth after incubation. Any zone of inhibition occurring around the disc is then measured, and after comparison with known standards, the bacterium under test is identified as susceptible or resistant to that particular antibiotic. For novel agents, these sensitivity parameters are only available after extensive clinical investigations are correlated with laboratory generated data. Disc tests are basically qualitative; however, the diameter of the zone of inhibition may be correlated to MIC determination through a linear regression analysis (Figure 18.6).


Although subtle variations of the disc test are used in some countries, the basic principles behind the tests remain similar and are based on the original work of Bauer and colleagues (Kirby–Bauer method). Some techniques employ a control bacterial isolate on each plate so that comparisons between zone sizes around the test and control bacterium can be ascertained (i.e. a disc potency control). Provided that discs are maintained and handled as recommended by the manufacturer, the value of such controls becomes debatable and probably unnecessary. Control strains of bacteria are available which should have inhibition zones of a given diameter with stipulated antimicrobial discs. Use of such controls endorses the suitability of the methods (e.g. medium, inoculum density, incubation conditions) employed. For slow growing microorganisms, the incubation period can be extended. Problems arise with disc tests where the inoculum density is inappropriate (e.g. too low, resulting in an indistinct edge to the inhibition zone following incubation), or where the edge is obscured by the sporadic growth of cells within the inhibition zone, i.e. the initial inoculum although pure contains cells expressing varying levels of susceptibility—so-called heterogeneity. As the distance from the disc increases, there is a logarithmic reduction in the antimicrobial concentration; the result is that small differences in zone diameter with antimicrobials (e.g. vancomycin) which diffuse poorly through solid media may represent significantly different MICs. Possible synergistic or antagonistic combinations of antimicrobials can often be detected using disc tests (Figure 18.4).


      ii)     Dilution tests


These usually employ liquid media but can be modified to involve solid media. Doubling dilutions, usually in the range 0.008–256 mg/L of the antimicrobial under test, are prepared in a suitable broth medium, and a volume of log-phase cells is added to each dilution to result in a final cell density of around 5 × 105 CFU/ml. After incubation at 35°C for 18 hours, the concentration of antimicrobial contained in the first clear tube is read as the MIC. Needless to say, dilution tests require a number of controls, e.g. sterility control, growth control, and the simultaneous testing of a bacterial strain with known MIC to show that the dilution series is correct. Endpoints with dilution tests are usually sharp and easily defined, although ‘skipped’ wells (inhibition in a well with growth either side) and ‘trailing’ (a gradual reduction in growth over a series of wells) may be encountered. The latter is especially evident with antifungal tests (see below). Nowadays, the dilution test for established antimicrobials has been simplified by the commercial availability of 96well micro titre plates which have appropriate antimicrobial dilutions frozen or lyophilized onto wells in the plate. The appropriate antibacterial suspension (in 200–400 μl volumes) is simply added to each well, the plate is incubated as before, and the MIC is read.


Dilution tests can also be carried out using a series of agar plates containing known antimicrobial concentrations. Appropriate bacterial suspensions are inoculated on to each plate and the presence or absence of growth is recorded after suitable incubation. Most clinical laboratories now employ agar dilution breakpoint testing methods. These are essentially truncated agar dilution MIC tests employing only a small range of antimicrobial concentrations around the critical susceptible/resistant cut-off levels. Many automated identification and sensitivity testing machines now use a liquid (broth) variant of the agar breakpoint procedure. Similar breakpoint antimicrobial concentrations are used with the presence or absence of growth being recorded by some automated procedure (e.g. light-scattering, colour change) after a suitable incubation period.


     iii)               E-tests


Perhaps the most convenient and presently accepted method of determining bacterial MICs, however, is the E (Epsilometer)-test. The concept and execution of the E-test is similar to the disc diffusion test except that a linear gradient of lyophilized antimicrobial in twofold dilutions on nylon carrier strips on one side are used instead of the filter-paper impregnated antimicrobial discs. On the other side of the nylon strip are a series of lines and figures denoting MIC values (Figure 18.5). The nylon strips are placed antimicrobial side down on the freshly prepared bacterial lawn and, after incubation, the MIC is determined by noting where the ellipsoid (pearshaped) inhibition zone crosses the strip (Figure 18.5).


For most microorganisms, there appears to be excellent correlation between dilution and E-test MIC results. As with standard disc diffusion tests, resistant strains may be isolated from within the zone of inhibition.


    iv)   Problematic bacteria


With some of the emerging antimicrobial-resistant bacterial pathogens, e.g. vancomycin-resistant enterococci (VRE), meticillin-resistant Staph. aureus (MRSA), vancomycin-intermediate Staph. aureus (VISA), the standard methodology described above may fail to detect the resistant phenotype. This is due to a variety of factors including heterogeneous expression of resistance (e.g. MRSA, VISA), poor agar diffusion of the antimicrobial (e.g. vancomycin) and slow growth of resistant cells (e.g. VISA). Disc tests are unsuitable for VRE, which should have MICs determined by E-test or dilution techniques. With MRSA, a heavier inoculum should be used in tests and 2–4% additional salt (NaCl) included in the medium with incubation for a full 48 hours. Reducing the incubation temperature to 30 °C may also facilitate detection of the true MIC value. Although 100% of MRSA cells may contain resistance genes, the phenotype may only be evident in a small percentage of cells under the usual conditions employed in sensitivity tests. Expression is enhanced at lower temperatures and at higher salt concentrations. With VISA, MIC determinations require incubation for a full 24 hours or more because of the slower growth rate of resistant cells.


b)  Tests For Bactericidal Activity


MBC testing is required for the evaluation of novel antimicrobials. The MBC is the lowest concentration (in mg/L) of antimicrobial that results in 99.9% or more killing of the bacterium under test. The 99.9% cut-off is an arbitrary in vitro value with 95% confidence limits that has uncertain clinical relevance. MBCs are determined by spreading 0.1 ml (100 μl) volumes of all clear (no growth) tubes from a dilution MIC test onto separate agar plates (residual antimicrobial in the 0.1 ml sample is ‘diluted’ out over the plate). After incubation at 35 °C overnight (or longer for slow-growing bacteria), the numbers of colonies growing on each plate are recorded.


The first concentration of drug that produces <50 colonies after subculture is considered the MBC. This is based on the fact that with MICs, the initial bacterial inoculum should result in about 5 × 105 CFU/ml. Inhibition, but not killing of this inoculum, should therefore result in the growth of 50 000 bacteria from the 0.1 ml sample. A 99.9% (3-log) kill would result in no more than 50 colonies on the subculture plate. With most modern antibacterial drugs, the concentration that inhibits growth is very close to the concentration that produces death, e.g. within one or two dilutions. In general, only MICs are determined for such drugs.

c)  Tests For Fungistatic And Fungicidal Activity


As fungi have become more prominent human pathogens, techniques for investigating the susceptibility of isolates to the growing number of antifungal agents have been developed. These have been largely based on the established bacterial techniques (disc, dilution, E-test) mentioned above, with the proviso that the medium used is different (e.g. use of RPMI 1640 plus 2% dextrose) and that the inoculum density (yeast cells or spores) used is reduced (c.104 CFU/ml). With yeast disc and E-tests, a lawn producing just separated/distinct colonies is preferable to confluent growth (see Figure 18.5). Addition of methylene blue (0.5 mg/ml) to media may improve the clarity of inhibition zone edges. Problems of ‘tailing’ or ‘trailing’ in dilution tests, and indistinct inhibition zone edges are often seen in tests involving azoles and yeasts and appear in some way related to the type of buffer employed in the growth medium. However, their presence has prompted studies into evaluating the use of other techniques as an indicator of significant fungistasis.


e.g. 50% reduction in growth (rather than complete inhibition) as the end point, use of a dye (e.g. Alamar blue) colour change to indicate growth, and sterol (ergosterol) quantitation. Most of these are presently outside the scope of most routine laboratories.


As with MBC estimations, MFC evaluation is an extension of the MIC test. At the completion of the MIC test (e.g. 72 hours for filamentous fungi), 20 ml are sub-cultured on to a suitable growth medium from each optically clear microtitre tray well and the growth control well. These plates are then incubated at 35 °C until growth is evident on the growth control subculture (24–48 hours). The MFC is the lowest drug concentration showing no growth or fewer than three colonies per plate to obtain approximately 99–99.5% killing activity.

d)  Evaluation  Of  Possible  Synergistic Antimicrobial  Combinations


The potential interaction between two antimicrobials can be demonstrated using a variety of laboratory procedures, e.g. ‘chequerboard’ MIC assays where the microorganism is exposed to varying dilutions of each drug alone and in combination, disc diffusion tests (see Figure 18.4) and kinetic kill curve assays. With the former, results can be plotted in the form of a figure called an isobologram (see Figure 18.7).



             i)  Kinetic Kill Curves


In the case of kill curves, the microorganism is inoculated into tubes containing a single concentration of each antimicrobial alone, the same concentrations of each antimicrobial in combination, and no antimicrobial i.e. four tubes. All tubes are then incubated and viable counts are performed at regular intervals on each system. With results plotted on semilogarithmic paper, synergy is defined as a greater than 100-fold increase in killing of the combination compared with either drug alone. Antagonism is defined as at least a 100-fold decrease in killing of the combination when compared with the most active agent alone, while an additive or autonomous combined effect results in a less than 10-fold change from that seen with the most active single drug. Both chemotherapeutic agents and disinfectants are amenable to kill curve assays.

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