2. Materials and Methods

Materials and Methods

2.1. Materials

  • The basic medium used in the CDS Test for the majority of organisms is Sensitest Agar (Oxoid CM409). Fastidious organisms will require the use of an enriched medium: Sensitest agar with 5% horse blood; Supplemented Haemophilus Test Medium Base (Oxoid CM898); Chocolate Columbia Blood Agar (Oxoid CM331); Supplemented Brucella Medium Base (Oxoid CM0169). Organisms requiring enriched media are specified in chapters 4, 5, 6, 7, 9 and 11.
  • “90 mm diameter” plastic Petri dishes.
  • Tubes with 2.5 mL (± 5%) of sterile isotonic saline.
  • 10 cm of 0.574 mm diameter nichrome wire (gauge B&S 23 or AWG 23) in a loop holder. The wire tip must be cut or filed square and must be replaced if it becomes worn or distorted. Available in lengths of one or more meters from:

Flame free laboratories can use disposable plastic inoculating needles from Copan Innovations (Supplied within Australia by Interpath Services – Catalogue No: 176CS20, Phone 1800 626 369 or 03 9457 6277). These needles yield a slightly heavier, but acceptable, suspension.1

  • Pasteur pipettes.
  • 6 mm diameter antibiotic discs supplied by Oxoid Pty Ltd (Thermo Fisher Scientific Australia Pty Ltd), Mast Group Ltd or other specified sources.
  • Disc dispenser (maximum of 6 discs) available from Oxoid Pty Ltd or Mast Group Ltd.
  • Max/min thermometer.
  • Clear plastic ruler, marked in millimetres.
  • 5 and 2 McFarland standards available from BD (Becton Dickson) and Thermo Fisher Scientific Australia Pty Ltd.
  • Spectrophotometer (optional).

 2.2. Methods

The nine steps followed in performing the CDS Test are represented diagrammatically in Figure 2.1. Further details of particular aspects of the method, including preparations necessary before the performance of the actual test, are set out below:

Figure 2.1 Performance of the CDS Test.

2.2.1. Agar plates


Prepare and handle dehydrated media strictly according to the manufacturer’s instructions.

Sensitest Agar: Oxoid Sensitest agar (CM409).

Blood Sensitest Agar: Prepare and sterilise Oxoid Sensitest agar (CM409). Cool to 50°C and add defibrinated horse blood to a final concentration of 5% (e.g. 100 mL defibrinated horse blood to 2 L prepared agar).

Haemophilus Test Medium: Prepare and sterilise Haemophilus Test Medium base (Oxoid CM898). Cool to 50°C and add fresh or deep frozen solutions of haematin and nicotinamide adenine dinucleotide (NAD) to a final concentration of 15 mg/L of each. Note: In-house prepared HTM must be used within 2 weeks of preparation.

Chocolate Columbia Blood Agar: Prepare and sterilise Columbia Blood Agar Base (Oxoid CM331). Cool to 70°C and add defibrinated horse blood to a final concentration of 9% (e.g. 180 mL defibrinated horse blood to 2 L prepared agar base). Hold at 70°C for 30 min to allow blood to chocolate. Cool to 50°C before pouring.

Supplemented Brucella Medium Base: Prepare and sterilise Brucella Medium Base (Oxoid CM0169). Cool to 50°C and add defibrinated horse blood to a final concentration of 5%, a fresh or deep frozen solution of haemin to a final concentration of 5 mg/L, and vitamin K to a final concentration of 1 mg/L.

            Alternatively, small laboratories can purchase Brucella supplemented agar plate (PP2459, Oxoid) to test anaerobes.

Plate preparation and storage

  1. Dispense 20 mL of agar into “90 mm diameter” Petri dishes.
  2. Store agar plates at 2‑8°C for a maximum of 4 weeks in sealed containers, plastic bags or shrink‑wrapped.
  3. Prior to use, surface dry the plates face down with the lid removed in an incubator at 35‑37°C. This will take approximately 1 hour in a fan‑forced incubator or 2 hours in an ordinary incubator.
  4. Store dried plates at 4°C and use within 2 days of drying.

NOTE: If commercial preparations that comply with the specification of the above media are used, the manufacturer’s instructions must be followed in all respects.

2.2.2. Preparation of the inoculum

The preferred method

  1. Use an overnight culture, preferably grown on blood agar, to prepare the CDS inoculum of 107 cfu/mL. With the straight wire, stab 1 colony (1 to 2 mm in diameter). Bacterial material should be visible on the tip of the straight wire.
  2. Inoculate the saline by rotating the straight wire at least 10 times with the tip in contact with the bottom of the tube.
  3. Mix up and down at least 10 times using a Pasteur pipette. With practice you should be able to see a faint turbidity in the saline suspension after mixing.

Using a Spectrophotometer

Prepare a suspension in 0.9% saline to achieve an absorbance of 0.15 at 640 nm (0.5 McFarland Standard). Dilute the suspension 1 in 5 (1 part suspension, 4 parts saline) in normal saline to obtain the CDS suspension.

Disposable plastic inoculating needles

The CDS method of antimicrobial susceptibility testing stipulates that a nichrome wire with a diameter of 0.574 mm (B&S 23) be used to collect the inoculum and create the CDS suspension. Open flames and other heat sources for sterilisation of metal inoculating needles and loops are not always available in microbiology laboratories. Gas may not be available in small laboratories, and open flames may be a fire hazard and a source of particulate pollution, which may have consequences for occupational health and safety. Other methods of sterilisation have similar disadvantages. Disposable plastic inoculating loops are widely used in microbiology laboratories for convenience and/or safety. Sterile disposable plastic inoculating needles (regular size) from Copan Innovation™ were compared with the recommended nichrome wire1. The Copan inoculating needle has a protruding cylindrical tip with a diameter of 0.64 mm. The Copan needles, in general, produced a marginally higher inoculum than the wire, but the CDS method was sufficiently robust to accommodate this increase and no major discrepancies in zone sizes were found.

Alternative methods for difficult organisms

With some organisms, using the preferred method may not yield visible bacterial material on the tip of the wire. In these situations one of the following acceptable alternative methods can be used.

Small colonies

Stab 3‑5 colonies (suitable for small colonies such as streptococci, haemophilus etc.).

Sticky colonies

Tease the colony apart and pick up bacterial material.

Tiny or pinHEAD colonies

Holding the straight wire at an angle of approximately 45°, move it in one direction along the edge of confluent growth until cellular material is just visible on the tip of the wire.

This is the least desirable method as the resulting inoculum may not be pure, but it may be necessary with Streptococcus anginosus (formerly S. milleri) and Streptococcus pneumoniae.

Scanty small / tiny colonies

In situations where very few colonies of Streptococcus pneumoniae have grown on the primary culture media, CDS sensitivity testing can be performed by growing the pneumococcal suspension in a tube containing 3 mL peptone water (10 g of peptone plus 5g sodium chloride in 1 L). To obtain the CDS inoculum of 107 cfu/mL, suspend 3 colonies (1 mm in diameter) or 6 colonies (0.5 mm in diameter) or 8 colonies (< 0.5 mm in diameter) in 3 mL peptone water and incubate at 35°C for 4 hours. The turbidity of the bacterial suspension should be visible to the naked eye.

Inoculum preparation for specific organisms


The inoculum is prepared from a pure culture of the organism grown for 24‑48 hours in an anaerobic atmosphere on any agar medium that will support its growth. Organisms are harvested from the surface of the agar and a bacterial suspension is prepared in 0.9% saline. The turbidity of the suspension is adjusted to an equivalent 0.5 McFarland standard. Alternatively the suspension can be adjusted to an absorbance of 0.15 using a spectrophotometer set at a wavelength of 640 nm.

Helicobacter pylori

The inoculum is prepared in Brain Heart Infusion broth (NOT SALINE) using a 48 – 72 hour culture of Helicobacter pylori grown on blood agar or chocolate agar (chocolate Columbia blood agar) in a microaerophilic atmosphere at 35‑37°C. The bacterial suspension should be adjusted to an equivalent 2.0 McFarland standard. Also, the suspension can be prepared by adjusting the absorbance to 0.3 using a spectrophotometer set at a wavelength of 640 nm. The inoculum gives a lawn of confluent growth (not semi‑confluent).


The inoculum is prepared using an 18 to 24 hour culture from a suitable Neisseria growth medium. Suspend a 1.0 to 2.0 mm colony in 2.5 ml saline. If colonies are small (<1.0 mm in diameter) suspend 3 to 5 colonies.

2.2.3. Inoculation of plates and application of discs

  1. Flood agar plate, rock the plate to distribute the suspension and remove excess with a Pasteur pipette.
  2. Remove the lid and place the plate, uncovered, on the bench to dry. This will usually take 5 to 10 min. Plates must not be left longer than 30 min.
  3. Apply no more than 6 antibiotic discs usually. In some circumstances an extra disc may be applied to aid in the phenotypic demonstration of particular phenomena such as in case of EDTA for confirmation of MBL, boronic acid for confirmation of AmpC and positioning of erythromycin and clindamycin for the detection of inducible clindamycin resistance.

NOTE: When testing Helicobacter pylori, apply only 3 discs per plate. With anaerobes, the inhibitory zone sizes around ticarcillin-clavulanic acid, meropenem and metronidazole might be very large and interfere with the reading of the inhibitory zones around neighbouring antibiotic discs. If necessary, repeat the test with fewer discs.

See Chapter 11 (Tables) for correct disc potencies and Section 3.3 (CDS‑QANTAS checklist) for correct storage and handling of stock and in use antibiotic discs.

2.2.4. Incubation of plates

Plates are placed immediately in the incubator so that there is no prediffusion and incubated upside down to prevent dehydration of the agar. Most susceptibility tests are performed either on Sensitest agar at 35‑37°C in air overnight, or on blood Sensitest agar at 35‑37°C in 5% CO2 overnight. However there are a few exceptions:

Anaerobes: Supplemented Brucella medium base at 35‑37°C anaerobically for 24 hours. Slow growing organisms will require 48 hours of incubation.

Campylobacter species: Blood Sensitest agar at 42°C in microaerophilic conditions.

Haemophilus species: HTM agar at 35‑37°C in 5% CO2

Helicobacter pylori: Chocolate Columbia blood agar at 35‑37°C in microaerophilic conditions for 72 hours.

Neisseria gonorrhoeae: Chocolate Columbia blood agar at 35‑37°C in 5% CO2 and > 80% humidity.

Yersinia enterocolitica: Sensitest agar at 30°C in air.

2.2.5. Organisms with special growth requirements

Cysteine, thymidine or glutamine requiring strains of Enterobacteriaceae and pyridoxal requiring streptococci (named Abiotrophia defectiva and Granulicatella adiacens) can be tested by adding 5 drops of a sterile aqueous solution containing one of the following: cysteine (2000 mg/L), thymidine (5000 mg/L), glutamine (1000 mg/L) or pyridoxal (1000 mg/L) to 2.5 mL of saline before inoculation. Sulphonamide and trimethoprim cannot be tested in the presence of thymidine.

CO2 dependent staphylococci can be incubated in 5% CO2 at 35‑37°C. The effect of CO2 on the zone sizes is not sufficient to influence the susceptibility test results. Some staphylococci grow poorly on Sensitest agar. Two notable examples include Staphylococcus lugdunensis and Staphylococcus pseudintermedius. In such cases, susceptibility testing should be performed on blood Sensitest. Reference ranges are provided in Table 11.3.a

2.2.6. Reading the zones

  1. Measure the zones from the back of the plate where possible.
  2. Measure the annular radius (the shortest distance from the edge of the disc to the edge of confluent growth). This usually corresponds to the sharpest edge of the zone (Figure 2).

Figure 2.2 Diagram showing the annular radius of the zone of inhibition.

2.3. Interpretation of results

See Chapter 11 (Tables) for the MIC breakpoints and the annular radius cut-offs for susceptibility to calibrated antibiotics, Surrogate antibiotics for antibiotics not calibrated, and specifics for the testing and reporting of β‑lactam antibiotics.

Standard Interpretation:

Annular radius            ≥ 6 mm            =          SUSCEPTIBLE

    < 6 mm            =          RESISTANT

Exceptions to the Standard Interpretation:

Exceptions to the standard 6 mm annular radius cut off are flagged in Tables 11.1 Calibrations.

2.3.1. Assessment of inhibitory zone morphology

In the CDS test we have adopted the practice of examining the morphology of the inhibitory zone when this may enhance the assessment of an organism’s susceptibility. The very oldest example of this is the observation of the sharp edge of the penicillin inhibitory zone with lactamase producing Staphylococcus aureus. Throughout the manual attention is drawn to those circumstances where assessment of susceptibility is enhanced by observing difference in zone morphology between susceptible and resistant isolates. In many cases, also, useful changes in zone morphology may be induced by the presence of an adjacent antibiotic disc so that disc placement becomes an essential component of the CDS test.


1    Outhred, A., Bell, S., Pham, J., Varettas, K., Rafferty, D. 2006. Performance of a disposable plastic inoculating needle in the preparation of the CDS inoculum. Annual Conference of