Microbiology Techniques manual

School of Biomedical Science Microbiology Techniques Manual Version 5.0 June 2002 ii TABLE OF CONTENTS Miscellaneous Section Laboratory Rules.......................................................................................................................1 Waste Disposal..........................................................................................................................1 Microscope Rules......................................................................................................................2 Setting Up The Microscope ......................................................................................................3 Decimal Dilution Series ............................................................................................................5 KOH String Test .......................................................................................................................6 Haemolysis................................................................................................................................7 Lawn Plate.................................................................................................................................8 McFarland Turbidity Standards ................................................................................................9 Preparation of a Bacterial Film for Staining ...........................................................................10 Spread Plate.............................................................................................................................11 Streak Plate .............................................................................................................................12 Total Bacterial Counts ............................................................................................................13 Identification Tests Aesculin/Bile Aesculin ...........................................................................................................15 Anaerobic Identification Discs................................................................................................16 Bacitracin Sensitivity ..............................................................................................................17 Bile Solubility .........................................................................................................................18 CAMP Test .............................................................................................................................19 Carbohydrate Fermentation—Acid Production ......................................................................20 Catalase ...................................................................................................................................20 Citrate......................................................................................................................................21 Coagulase—Slide....................................................................................................................22 Coagulase—Tube....................................................................................................................23 Composite Media ....................................................................................................................24 Decarboxylase Tests—Arginine, Lysine and Ornithine .........................................................26 DNAse.....................................................................................................................................27 Gelatin Hydrolysis ..................................................................................................................28 Germ Tube ..............................................................................................................................29 Hippurate Hydrolysis ..............................................................................................................30 Hydrolysis of Complex Substrates—Casein, Chitin, Tyrosine and Xanthine ........................31 Indole ......................................................................................................................................32 Methyl Red..............................................................................................................................33 Motility ...................................................................................................................................34 Neisseria Carbohydrate Fermentation.....................................................................................36 i Nitrate Reduction ................................................................................................................... 37 Novobiocin Sensitivity ........................................................................................................... 39 O/129 Sensitivity.................................................................................................................... 40 ONPG (β-Galactosidase)........................................................................................................ 40 Optochin Sensitivity ............................................................................................................... 41 Oxidase................................................................................................................................... 42 Oxidation—Fermentation (O-F) Test..................................................................................... 43 PYR Test ................................................................................................................................ 44 Salt Tolerance......................................................................................................................... 44 SPS Sensitivity ....................................................................................................................... 45 Superoxol ............................................................................................................................... 46 Triple Sugar Iron .................................................................................................................... 47 Urease..................................................................................................................................... 48 VP (Voges-Proskauer)............................................................................................................ 48 X and V Factor Requirement.................................................................................................. 49 Antibiotic Sensitivity Testing Calibrated Dichotomous Sensitivity (CDS) Method .............................................................. 51 Disc Diffusion NCCLS (Kirby-Bauer) Method ..................................................................... 53 Staining Albert’s Stain—Volutin Granules .......................................................................................... 55 Capsule and Negative Stain.................................................................................................... 55 Flagella Stain.......................................................................................................................... 56 Gram Stain.............................................................................................................................. 57 Kinyoun Stain......................................................................................................................... 58 Lactophenol Cotton Blue Tape Preparation ........................................................................... 59 Methylene Blue (Simple) Stain .............................................................................................. 59 Spore Stain ............................................................................................................................. 60 Ziehl-Neelsen (ZN) Stain ....................................................................................................... 61 ii LABORATORY RULES THESE RULES ARE EXTRACTED FROM THE FULL MICROBIOLOGY SAFETY MANUAL AVAILABLE AS HARDCOPY IN THE PREPARATION ROOM, OR ON LINE AT http://www.healthsci.utas.edu.au/biomed/units/micro/safety.html 1. 2. 3. 4. Coats, jackets and other outer apparel should be left outside the laboratory, together with bags and books not required for the laboratory session. Long hair should be tied back neatly, away from the shoulders and enclosed footwear should be worn—(thongs and open sandals are not allowed). Avoid placing any object in your mouth—(pencils, pens, fingers etc). Mouth pipetting is strictly forbidden in the microbiology laboratory. Laboratory gowns are provided for your protection and must be worn inside the laboratory. They are not to be worn outside the laboratory for any reason. At the end of the laboratory session please return the gown to the hook, neatly folded, inside out. No slides or cultures are to be taken from the laboratory. If you have an accident of any kind call the instructor immediately. A spilled culture can be covered with paper towels and laboratory disinfectant poured over the towels and contaminated area; leave for 15 minutes before mopping up. The working area should be wiped with disinfectant at the beginning and end of the laboratory session. Always wash your hands before leaving the laboratory. 5. 6. 7. 8. WASTE DISPOSAL The waste disposal protocol in the Microbiology Laboratory is designed to separate the non-infectious from the infectious waste. The infectious waste needs to be disposed of in a manner that minimises the risk to both staff and students and facilitates the recycling of reusable material. Please follow the instructions carefully and if in doubt—ASK! Sharps One dedicated yellow sharps container per laboratory. Used for needles, scalpel blades and infectious broken glass. Do not wander around the laboratory carrying sharps, ALWAYS take the container to the sharps, NOT the sharps to the container! Biogram buckets Located on each bench. Used for contaminated waste, eg used swabs, capillary tubes, pipettes and wet slide preparations. Not to be used for gram stains, paper, matches or chemicals. Biohazard bin Located in the middle of the laboratory. Used for contaminated waste, eg used culture plates and contaminated paper towel. Not to be used for paper or glass. Billy cans Two of these are located at the front of the laboratory. 1 1. 2. Used for recyclable glass or plastic tubes/bottles. Please remove any sticky labels from these. Used for fixed and stained slides (not for wet preps). Paper bin For non contaminated paper only, eg. paper towel from hand washing or blotting gram stains. MICROSCOPE RULES When you have finished using your microscope 1. 2. 3. Remove slides from the stage. Turn light source down to 0. Clean objectives if using oil. 2 4. 5. 6. 7. Switch off. Leave lowest power objective (ie x4) in viewing position. Put cover on. Turn off at the power point. 3 SETTING UP THE MICROSCOPE 1. Before use check for: a) fingerprints, immersion oil, dust, etc. on objective front lens, eyepiece surfaces, condenser top lens, and light exit glass on microscope base. All fingerprints and other stains should be carefully removed with lens tissue moistened in Gram’s acetone-alcohol; fine dust on all parts of the microscope. Any present should be gently removed with an air blower or by using a camel hair brush. WHEN CLEANING LENSES PLEASE USE ONLY LENS TISSUE b) 2. Microscopes provided are either Olympus, Nikon or Industrial Scientific binocular a) b) c) d) e) Check that the power dial is set to low. Turn on at the power point. If the microscope is fitted with a phase contrast ring turn it so that the number 0 can be seen. Switch on light source by means of the ON-OFF button on microscope base. Place an object slide on the stage and adjust the light intensity by the brightness adjusting dial or knob which is also on the microscope base. The power should be adjusted to about 50% of the maximum or so that viewing is comfortable. Using the x10 objective and the coarse focus, bring the object on the slide into focus. Adjust the interpupillary distance by moving the eyepieces in or out until a single circular image can be seen. Now adjust the focus for differences in your eyes. Check your eyepieces to determine which one has a dioptre adjustment ring, which when turned will move the eyepiece in or out. Look through the eyepiece which does not have a dioptre adjustment ring and bring the specimen into focus with the coarse and fine adjustment knobs on the microscope stand. Use the 10x objective. Now look through your opposite eye and adjust the focus using the focus ring on the eyepiece. This will compensate for small differences in focal length between your eyes. Adjust the microscope for brightfield viewing; f) g) h) Nikon Microscopes Fully close the diaphragm which is on the illuminator on the base of the microscope. Focus the light source by moving the position of the condenser up and down (not the objective lens) while looking through the eyepieces. You should see the light rays refract as they pass the diaphragm. This will cause the light rays to diverge according to their wavelength and you will see different colours of light. You will also see the edges of the diaphragm move in and out of focus. The correct position of the condenser is achieved when the edges of the diaphragm are sharply focused. Centre the condenser using the centring screws so that the diaphragm appears to be in the middle of the viewing area. Open the diaphragm until the edges no longer impinge upon the viewing area. These microscopes are more basic in design than the Nikons and must be set up slightly differently. Focus the light source by racking the condenser (not the objective lens) fully up and then back it off approximately 2-3 mm. You may close the diaphragm on the condenser to about 70-80% of the objective numerical aperture. This will improve image contrast and increase depth of focus. Olympus and Industrial Scientific Microscopes 4 3. Fine focussing on the specimen a) Commence with the lowest power objective (X4 or X10) and use the coarse focusing knob. Once the image is sharply focused swing in the next objective and repeat the procedure. Once you start viewing with the X40 objective use only the fine focusing ring. A major problem that many inexperienced students have is a poor image when using the X40 objective. Often this is a result of a dirty objective. If this happens to you clean the front element of the objective with lens tissue. When looking at stained preparations of bacteria you will need to view them with the X100 objective which requires immersion oil to be placed between the specimen and the objective. Tilt the X40 objective to the side, place a small drop of oil on the slide and swing in the oil immersion lens (X100) to make contact with the oil. The objective may not be parfocal, so focus again using a slight clockwise movement of the fine adjustment. NEVER adjust the coarse focus when using the high power objectives, x40 and x100. On completion of oil immersion observation swing out oil immersion objective before removing slide. ALWAYS ROTATE THE NOSEPIECE SO THAT THE X40 OBJECTIVE NEVER PASSES OVER A SLIDE WITH OIL ON THE SURFACE. THE OIL IMMERSION (X100) HAS A BLACK RING. PLEASE BECOME FAMILIAR WITH THIS AND ENSURE THAT ONLY THE X100 OBJECTIVE IS INSERTED INTO THE OIL. 5 DECIMAL DILUTION SERIES Often for experiments or when quantifying bacteria a serial dilution is necessary before culturing. When counting live organisms by culture we aim to achieve between 30-300 colonies on a plate for statistical accuracy. When dealing with unknown amounts of cells, several dilutions should be cultured to ensure this range of colonies can be counted. Reagents and Materials 4.5 ml or 2.5 ml diluents (use same throughout, example below uses 4.5 ml) Sterile 1.0 ml pipettes Method 1. Take a sterile 1 ml pipette and attach rubber teat. Suck the neat suspension up and transfer 0.5 ml to 4.5 ml diluent (10-1 dilution). Discard the pipette into disinfectant. Take a fresh pipette suck up the 10-1 dilution and down ten times (alternatively mix the suspension on a vortex) and transfer 0.5 ml from the 10-1 dilution to the next tube (10-2 dilution). 2. 3. Repeat the procedure to the required dilution. Suspension for dilution 4.5 mL 4.5 mL 4.5 mL 4.5 mL Dilution Decimal dilution Label Dilution factor (for calculation) 1 in 10 10-1 -1 101 1 in 100 10-2 -2 102 1 in 1000 10-3 -3 103 1 in 10000 10-4 -4 104 6 KOH STRING TEST The Gram stain reaction is not always truly indicative of the organisms true cell wall structure. Poorly controlled decolourising can obviously be a source of misleading results, but even with adequate technique some organisms are problematic. For example, some strains of Bacillus or Clostridium consistently stain Gram negative even though they have a Gram positive type cell wall. Reagents and Materials 3% Potassium hydroxide Test Procedure Emulsify a loopful of colony and a drop of 3% KOH on a glass slide. Stir the suspension continuously for 60 seconds and then gently pull the loop vertically away from the suspension. Interpretation Gram negative bacteria produce a viscous mixture and a thread of this viscid material follows the loop for 10 mm or more as it is raised. Controls Positive Negative Any known Gram negative organism Any known Gram positive organism 7 HAEMOLYSIS When bacteria are grown on blood agar, many produce extracellular products (haemolysins) that lyse or alter the pigment in the red cells; this is termed haemolysis. Since blood agar is one of the most widely used primary culture media, the presence and type of haemolysis has become a very useful criteria for identification. Type of haemolysis forms the basis for the classification of the clinically significant streptococci and is very important in many other organisms, both in terms of recognition of organisms in primary culture and subsequent identification. Alpha Haemolysis Characterised by an indistinct zone of partially destroyed red cells around the colony; the cells usually display a brown-green colouration due to alteration of the normally red haemoglobin pigment. Beta Haemolysis Characterised by a distinct clear colourless zone around the colony caused by the total destruction of the red cells. Gamma Haemolysis A ridiculous term used to describe the absence of haemolysis. Really only used in American textbooks. Use the term at your own risk. Partial Haemolysis Characterised by an indistinct zone of partially destroyed red cells around the colony. In contrast to alpha haemolysis, the haemoglobin pigment remains red but due to partial destruction the blood appears a lighter shade of red compared to areas without haemolysis. Double Zone Haemolysis This is the term used to describe zones of partial haemolysis when they are found around an inner zone of beta haemolysis. Types of Haemolysis Detecting Haemolysis Not all zones of haemolysis are obvious and students often fail to detect small or indistinct zones. As with most things in life, more satisfactory results can be obtained if one “looks toward the light!”. To do this remove the lid from the plate and hold the plate—with the agar surface facing you—towards a window or other light source. Type of Blood In many cases, haemolysins are specific to red cells from a particular animal—so a particular organism may be beta haemolytic on sheep blood agar but not on horse blood agar. Most Australian and European laboratories use horse blood, while American laboratories use sheep blood. This can be confusing since most text books are produced in America and describe haemolysis in terms of sheep blood. The plates at this campus of the University of Tasmania are prepared with sheep blood – unless otherwise indicated. Atmosphere of Incubation Some organisms show different haemolytic properties depending on the atmosphere in which the plates are incubated. This is classically so for some strains of Streptococcus pyogenes that produce an oxygen labile haemolysin and are beta haemolytic under anaerobic conditions but non haemolytic when incubated aerobically. Presence of Other Organisms In mixed cultures it is not uncommon to see extracellular bacterial products that are non haemolytic alone, combine to cause haemolysis or to combine to cause a different type of haemolysis. This property is used in some diagnostic tests (see CAMP test). Factors Affecting Haemolysis Describing Haemolysis When you are describing the haemolytic properties it is important that the conditions described above are considered. Always state the source of the red cells and the conditions of incubation. It is also important to describe the haemolysis in terms of the size of the zone. It is usually convenient to describe the size in relation to the size of the colony. 8 LAWN PLATE Lawn plates provide confluent even growth of a pure culture over an entire plate. They are most often used for sensitivity testing, where a disc or discs are placed on the plate to test the organism against certain substances. They can also be used to harvest a very large amount of organism for further testing. Reagents cotton swab agar plate sterile saline or other diluent. Inoculum The inoculum may already be in liquid form, eg. a urine specimen or a broth culture. If you have a culture on a solid medium, transfer a portion of a colony (or more if needed) to a tube of sterile diluent. Saline is usually used but more sophisticated diluents may be necessary in some cases; water should be avoided. Mix the organism and diluent to give a smooth suspension. The turbidity of the suspension can be standardised by comparison with McFarland Standards if required (see method). The inoculum can be applied to the plate in one of two ways. Swab Method Dip a sterile swab into the inoculum and lightly rotate the swab. Remove the excess fluid by rotating the swab against the inside of the tube above the level of the liquid. Cover the entire plate by swabbing in three directions (see fig. 1). It is important that the plate is covered evenly—so make each stroke with the swab touch the previous stroke and avoid a series of discrete individual lines of inoculation. Fig 1. Lawn Plate Step 1 Step 2 Step 3 Pour Method This method is more time consuming but provides a more consistent lawn plate. Use predried plates—plates can be dried by placing in the 37˚C incubator with lids off for 30 mins. Pour approximately 2.5 ml of the inoculum onto the plate and rotate the plate to achieve total coverage. Remove the excess inoculum with a Pastuer pipette—this is easier if you tilt the plate slightly. Dry the plate by leaving at room temperature with the lid off for no more than 30 mins before applying any discs to the plate. 9 MCFARLAND TURBIDITY STANDARDS When making bacterial suspensions, it is sometimes desirable to know how many cells are present or sometimes necessary to make an inoculum up to a predetermined concentration. The larger the concentration of bacterial cells, the greater the turbidity of the resulting suspension. Various concentrations of a relatively stable suspension of barium chloride particles have been calibrated to have a similar turbidity to a range of bacterial concentrations. Reagents Set of McFarland Standards Sterile diluent Method Make a suspension of bacterial cells in a transparent, colourless diluent and compare the turbidity against the McFarland standards. Comparisons of turbidity are best made by examining printed words (or black lines) on a white background through the tube and suspension. It is important that the tubes used to suspend the bacterial cells in are as close in size shape and composition to those in which the standards are made—at the very least the width of the tubes must be the same. Interpretation McFarland Standard Bacterial cells x 10 8 per ml 0.5 1.5 1 3 2 6 3 9 4 12 5 15 6 18 7 21 8 24 9 27 10 30 Precautions The McFarland standards are relatively stable if kept in the dark but consideration should always be given to the age and reliability of the standards. The calibration is VERY approximate and a large number of other variables will affect the turbidity in addition to the concentration of bacterial cells. Some important variables to consider are the shape and size of the organism, the growth medium it was taken from and the phase of growth the organism was in when the suspension was prepared. Visual inaccuracies can be avoided by using a spectrophotometer. 10 PREPARATION OF A BACTERIAL FILM FOR STAINING Before staining the bacterial cells must be applied to the slide in a manner which facilitates their staining and viewing with a microscope. Important points to remember when preparing the film are; Do not make the film too thick since this will make it difficult to decolourise the cells and will make viewing individual cells difficult. Do not apply too much water to emulsify the cells with since it will take too long for the slide to dry before you can fix the cells to the slide. Film preparation requires a balance between using the correct amount of water and cells. Remember even if you can’t see the cells on the slide before staining they are still there. Materials Glass slides Diamond/Glass pencil Water Loops and loop rest Bacterial culture Forceps Method 1. 2. Choose a clean and dry slide. Label slide carefully using a glass or diamond pencil. Three films can be prepared on the one slide with a ring about 1 cm diameter for each as shown below. 1 3. 4. 5. 2 3 Sterilise the inoculating loops in the flame and allow them to cool on the loop rest Place a small volume of water or culture broth in the centre of each circle. This is best done with a loop. Touch the colony to be studied with the loop and emulsify a portion in the water drop, spreading it to cover the circled area. Do not take more than one half of one colony. Sterilise the loop and replace it on the loop rest; use the second loop to repeat the procedure for colony 2 and so on. Allow slides to dry at room temperature; After the smear is dry, fix bacteria to the slide by passing it through the flame with the film side uppermost. The total period of heating should not be more than one to two seconds. Hold the slide with forceps for this procedure. Heat fixing kills the bacteria and makes them adhere to the slide. Your slide is now ready for staining. 6. 7. 8. 11 SPREAD PLATE This method of culture can be used for the determination of viable cells in solution. It is relatively quick and easy but has a number of inaccuracies and requires numerous replicates for statistical accuracy. Reagents and Materials Diluted bacterial cells Plate count agar Pasteur pipettes Alcohol Volumetric pipettes Method 1. 2. 3. Prepare appropriate dilutions of the organism to be tested. Neatly label all petri dishes with the appropriate dilutions, experiment and date. Prepare several glass spreaders from pasteur pipettes. Fuse the end of the pipette in the bunsen flame and then heat to bend the pipette in two places according to the diagram. 4. 5. Sterilise a spreader by dipping it in alcohol, shaking off the excess alcohol and flaming. Allow to cool. Pipette 0.1 ml of the lowest dilution onto the surface of the appropriately labelled agar plate. Spread immediately as some cells are inclined to stick in situ. Try to spread as evenly as possible, giving a uniform coverage right to the edges. 6. 7. 8. Proceed with the remaining dilutions using the same pipette and a sterile cool spreader. Incubate the inverted plates at the appropriate temperature in a closed container. Count the colonies on the plate having between 30 and 300 (the lower limit is set by statistical accuracy and the upper limit by coincidence limitations). Observe the plates before the colonies become too large; this makes counting easier. 12 STREAK PLATE The streak plate is used to produce cultures containing well isolated single colonies that will demonstrate typical colonial morphology, can be used for further testing or can be subcultured to provide pure cultures. Materials Method Microbiological loop Bunsen Burner Agar plate Primary inoculum: Using a sterile loop pick up your inoculum—this may be a colony from another plate or some liquid from a broth culture—and transfer it to the fresh plate. Spread this evenly over about 1/4 of the plate as indicated in Fig. 1. This is the primary inoculum. The primary inoculum can also be made with a cotton swab—this is usual if the primary inoculum is actually a clinical specimen and useful if you are subculturing from a broth. Sterilise the loop by flaming. Streaking: With a sterile loop, make a series of strokes, starting with the loop in the primary inoculum and dragging the inoculum across the plate diluting it as you go. These strokes should be made in one direction only—NOT backwards and forwards. Repeat this process twice to give three sets of diluting streaks. Finally make a single streak that starts of in the end of the last set of diluting streaks and zig zags across the remainder of the plate. This final streak should produce isolated colonies—and it is very important that this streak does not come into contact with any of the other preliminary diluting streaks or the primary inoculum. Note: The loop should always be sterilised after the primary inoculum. With a heavy inoculum or for inexperienced students it is advisable to also sterilise the loop between each subsequent set of streaks. However, with a light inoculum or for more experienced students it may be possible to achieve satisfactory results without reflaming between streaks. To achieve this, think of the loop as having three surfaces—left edge, right edge and top edge (see Fig 2). Rotate the loop between sets and essentially use a fresh “sterile “ surface in each section of the plate. Fig 1 Streak Plate = primary inoculum Fig 2 Microbiology Loop edge 1 edge 2 edge 3 13 TOTAL BACTERIAL COUNTS The Improved Neubauer counting chamber has a total ruled area of 9 mm2 and a depth of 0.1 mm. The central area (1 mm2) is divided into 25 squares, each with an area of 0.04 mm2, and each of these are further marked into 16 squares. The volume of the diluent contained between the central square and the coverslip is 0.1 mm3 which is equivalent to 0.1 µL. Materials and Reagents Improved Neubauer counting chamber Coverslip Capillary Tube Bacterial suspension Phase contrast microscope Figure 1. Side view of a Neubauer counting chamber Figure 2. Ruled area of an improved Neubauer counting chamber. When counting bacterial cells, calculated a mean by using the squares marked with an R. 14 Method 1. 2. 3. 4. Check that the chamber and coverslip are clean and dry. Moisten the edges of the coverslip and slide it over the central ruled area of the counting chamber. Use a capillary tube to sample the bacterial suspension. Apply the capillary tube to the edge of the coverslip allowing the bacterial suspension to flow into the counting chamber. Do not overfill the counting chamber so that the bacterial suspension flows into the vee shaped edges. Stand the chamber for up to 20 minutes to let the cells settle onto the bottom, so that they are all in one focal plane. Examine under a microscope using the x40 objective and count the cells using squares in a predetermined order. From the area of the squares counted, the depth of the chamber and the original dilution factor, it is possible to arrive at a value for the total number of organisms per ml. 5. 5. Calculation Cell Count per litre = N x DF x 10 6 AxD N = average number of cells counted, DF = dilution factor, 106 = conversion factor for cells per litre, A = area counted and D = depth of chamber Size of one small square = 15 x 1 mm 5 Depth of counting chamber = 0.1 mm Therefore Cell Count per litre = N x DF x 10 6 1 x 1 x 0.1 5 5 N x DF x 106 = 1 x 0.1 25 ( ) N x DF x 107 = 1 25 = 25 x N x DF x 10 7 = 2.5 x N x DF x 10 8 Therefore Cell Count per ml = 2.5 x N x DF x 10 5 15 AESCULIN/BILE AESCULIN A test to determine the ability of an organism to hydrolyse aesculin either in the presence or absence of bile. The bile aesculin test is useful for the differentiation of Group D Streptococci and Enterococci from other Streptococci while hydrolysis in the absence of bile is useful for differentiation of Enterobacteriaceae. Reagents and Materials Inoculation Bile aesculin agar slope, plate or aesculin broth. Bile aesculin: using a straight wire or loop, lightly streak the surface of the agar slope with the organism to be tested. For bile aesculin plates, use spot inoculum and test multiple organisms per plate. Aesculin broth: lightly inoculate the broth with the organism to be tested. Incubation Streptococci: 48-72 hours Enterobacteriaceae: 18-24 hours Enterococci: 1 – 2 hours or less with reasonable inoculum Interpretation Aesculin hydrolysis is indicated by the formation of a black precipitate throughout the media or on the agar slope. Growth without aesculin hydrolysis may occur, and in the presence of bile indicates bile tolerance. The bile aesculin test is frequently used for presumptive identification of Enterococci. A positive result on a catalase negative gram positive coccus is presumptive identification of an enterococcus, although it should be noted that other less commonly isolated streptococci may also be positive. Controls: Bile Aesculin Positive Negative Enterococcus faecalis Streptococcus agalactiae(Group B streptococcus) 17 ANAEROBIC IDENTIFICATION DISCS The identification of the anaerobic Gram negative bacilli has not reached the level of specificity or convenience associated with the identification of their aerobic counterparts. However, the anaerobic Gram negative bacilli can be assigned to broad and clinically relevant groups based on sensitivity to a range of specific antimicrobial discs. Reagents and Materials Colistin disc (10µg) (CT DD) Vancomycin disc (5µg) (VA DD) Kanamycin disc (1000µg) (K DD) Erythromycin disc (60µg) (E DD) Penicillin disc (2 units) (P DD) Rifampicin disc (15µg) (RD DD) Blood agar Inoculation Prepare a lawn plate (see method) on blood agar (or other blood based enriched agar) using a 0.5 McFarland standard. Place the discs about 20 mm from the edge of the plate, equally spaced and in a circular arrangement. Incubation Incubate overnight, anaerobically @ 37˚C. Test Procedure Interpretation Measure the zones of inhibition. A zone daimeter > 10 mm is considered sensitive. The interpretation is based on the assumption that the organism is a Gram negative anaerobic bacillus. ORGANISM Bacteroides fragilis Prevotella sp Fusobacterium sp Porphyromonas sp R = resistant E S V R V S = sensitive R S V R V P R V S V V = variable VA R R R S K R R S R CT R V S R Caution It is prudent to also perform tests that confirm the anaerobic nature of the organism. This could be a metronidazole susceptibility test (where the MTZ5 disc is included on the plate with the ANIDENT discs), or an actual test for atmospheric requirements. 18 BACITRACIN SENSITIVITY Sensitivity to bacitracin is a presumptive test for the identification of Streptococcus pyogenes. The test can also be used to screen for the presence of the organisms on primary isolation plates. Reagents and Materials Diagnostic bacitracin disc (0.04 U) Blood agar or blood sensitest agar (if necessary, see inoculation and incubation) Inoculation and incubation The disc can be included with the therapeutic discs in a standard antimicrobial susceptibility test. In this case the plates should be incubated overnight in air @ 37˚C. Alternatively, the disc may be added to the blood plates used for primary isolation from clinical specimens, or on a simple subculture streak plate. The disc should be placed on the plate at the junction of the first and second streak lines (see OPTOCHIN SENSITIVITY TEST). In this case, incubation should be overnight in air, CO2 or anaerobically @ 37˚C. It is not necessary to measure the zone of inhibition; any zone of inhibition should be considered sensitive. Be careful to look for an actual zone of inhibition of growth—the surrounding zone of haemolysis may actually go up to the disc. Interpretation is based on the assumption that the organism has been confirmed as a beta-haemolytic streptococci. Bacitracin sensitive: presumptive Streptococcus pyogenes Bacitracin resistant: organism is NOT Streptococcus pyogenes Interpretation Controls SENSITIVE Streptococcus pyogenes (GpA) RESISTANT Streptococcus equi or equisimilis (GpC) 19 BILE SOLUBILITY This is a rapid and alternative test for the identification of Streptococcus pneumoniae: it is useful for those isolates with equivocal optochin sensitivity or for rapid presumptive identification of primary isolates. Reagents and Materials 2 x 2.5 ml salines 10% bile salts (Na desoxycholate or Na taurocholate) Inoculation Combine the 2 salines into a single tube and make a suspension of the organism equivalent to a 1.0 McFarland. Pour half the inoculum back into the empty tube: you should now have 2 identical tubes. Add 2-3 drops of bile salt solution to one tube (test) and 2-3 drops of saline to the other (control). Incubation Incubate @ 37˚C for 30 mins. Test Procedure Observe for bile solubility which is indicated by significantly decreased turbidity in the test tube compared to the control tube. Interpretation Controls Only Streptococcus pneumoniae is bile soluble. Positive Negative Streptococcus pneumoniae Enterococcus faecalis 20 CAMP TEST CAMP is an acronym from Christie, Atkins and Munch-Peterson, the Australians who devised a presumptive test for the identification of Group B streptococci based on the ability of that organism to enhance the zone of haemolysis around specific strains of Staphylococcus aureus. The term “CAMP TEST” has now come to include a range of similar tests where extracellular products from pairs of organisms interact to modify zones of haemolysis. Reagents and Materials Sheep blood agar (source of red cells is critical) ß-lysin producing Staphylococcus aureus (indicator organism) Inoculation A single, straight, horizontal streak is made across the centre of the plate with the indicator organism. A single straight vertical streak of the test organism is made upto—BUT NOT TOUCHING—the indicator organism. It is important that the two streak lines are very close, but do not touch. Overnight in air @ 37˚C, except reverse CAMP test with Clostridium perfringens which must be incubated anaerobiucazlly. Incubation Test Procedure Examine the area where the streak lines interact and look for alteration in haemolysis. With the “Reverse CAMP Test” (see below) the plates should be further incubated at 4˚C for 30 minutes immediately prior to examination. Interpretation Group B streptococci and Listeria monocytogenes both enhance the haemolysis of ß-lysin producing Staphylococcus aureus . A larger zone is produced with Group B streptococci. Corynebacterium pseudodiphtheriticum, Arcanobacterium haemolyticum and Corynebacterium ulcerans actually inhibit the haemolytic effect of ß-lysin producing Staphylococcus aureus . This is referred to as a “Reverse CAMP Test”. Another modification is also occasionally called the “Reverse CAMP Test”—not because haemolysis is inhibited—but because the Group B Streptococcus is used as the indicator organism and Clostridium perfringens is used as the test organism. With this modification, there is enhancement of haemolysis but no requirement for a 4˚C incubation period. There are other modifications—especially with respect to speciation of Listeria—but those listed above are the most commonly used. See Figure 1 21 CARBOHYDRATE FERMENTATION—ACID PRODUCTION A test to determine the ability of an organism to ferment a specific carbohydrate incorporated into a basal medium producing acid or acid with visible gas. Gas production is generally only tested with glucose since organisms which produce gas from this carbohydrate will also produce gas from other carbohydrates. Reagents and Materials Appropriate carbohydrate broth (usually a 1% concentration of carbohydrate in a peptone base with a pH indicator) Inoculation Incubation Transfer a portion of the colony to be tested to the media containing the appropriate carbohydrate. 18-24 hours. Incubation of up to 3 days may be required for some organisms. Check results daily. Interpretation Acid production is indicated by a change in colour of the indicator used. Bromocresol purple is most commonly used which is purple in alkali and yellow in acid. Gas production is indicated by displacement of liquid from the inverted Durham tube. Full Gas Incubation CATALASE A test for the presence of catalase enzyme present in most cytochrome containing aerobic and facultative anaerobic bacteria. Primarily used to differentiate between the genera Streptococcus from Micrococcus and Staphylococcus; Bacillus from Clostridium and Listeria monocytogenes and/or Corynebacterium from Erysipelothrix. Reagents and Materials Hydrogen peroxide 3% glass tube wooden tooth pick or glass rod Test Procedure Add 0.5-1.0 ml of reagent to a clean glass tube. Using a wooden tooth pick or a glass rod, transfer a portion of the colony to be tested to the tube. Care should be taken when using cultures from media containing blood since free catalase may be present. Positive and negative controls should always be used. Interpretation Rapid liberation of gas indicates catalase enzyme. Formation of static bubbles adhering to the applicator does not indicate a positive result. Controls Positive Staphylococcus spp. 22 Negative Streptococcus spp. 23 CITRATE (SIMMONS METHOD) Introduction Tests the ability of bacteria to utilise citrate as the sole source of carbon in media containing (NH2)2PO4 as the sole source of nitrogen. Organisms that cannot utilise citrate will not grow, while those that do grow, produce alkaline by-products and change the colour of the medium from green to blue. Most commonly used as one of a battery of tests in the characterisation of the Enterobacteriaceae. Reagents and Materials Inoculation Simmons citrate agar—as plate or slant. Use a straight wire to very lightly inoculate the surface of the medium. Heavy inocula may allow carry over carbon sources and give false positives. If a plate is being used, multiple organisms can be tested on the same plate provided there is sufficient space between organisms to prevent misinterpretation. Incubation 24—48 hours at 37˚C Interpretation Negative – Green Positive – Blue Some organisms may require extended incubation in order for the colour change to become apparent. An excellent test for the presumptive identification of E. coli from urine cultures. A citrate and oxidase negative, indole positive, lactose fermenting organism from a urine specimen is presumptively identified as E. coli. Controls Positive Negative Pseudomonas aeruginosa Escherichia coli 24 COAGULASE—SLIDE A presumptive and rapid test to assist in the identification of staphylococci. The slide coagulase is distinctly different from the tube coagulase and detects a surface protein on staphylococcal cells that cross links with fibrinogen in plasma and causes a clumping of bacterial cells. The test is often referred to as clumping factor. Reagents and Materials Glass slides Oxalated plasma (preferably rabbit but human plasma can be substituted) Distilled water or saline Inoculum This test can only be performed on colonies grown on non selective media. Test Procedure Add two loopfuls of water or saline to a glass slide. Add a portion of a colony to a spot on the slide adjacent to but not touching the diluent. With the loop, bring the diluent to the inoculum and mix until a smooth heavy milky suspension is obtained. If the suspension agglutinates at this stage, try changing diluent (water or saline) but if agglutination continues the isolate is said to be autoagglutinable and is unsuitable for the slide coagulase test. Add a slightly larger volume of plasma to a spot on the slide adjacent to but not touching the smooth suspension. Use the loop to bring the suspension into the plasma and mix. Visible clumping within 10 secs is a positive test. Interpretation This test lacks the specificity of the tube coagulase. A positive test is very suggestive and a negative test almost exclusive of S. aureus. The test should not be used in isolation but in conjunction with other information such as colonial morphology—or preferably with other presumptive tests such as mannitol fermentation or DNase production. Controls Positive Negative Staphylococcus aureus Staphylococcus epidermidis Caution The most common problem that students have with this test is due to the suspension of organism being too weak. Follow the method exactly as described and MAKE SURE YOU HAVE A SMOOTH HEAVY MILKY SUSPENSION. 25 COAGULASE—TUBE The definitive test in the identification of Staphylococcus aureus. The test detects the presence of the free coagulase enzyme, which has a thrombin like action and is able to clot plasma. Reagents and Materials Trypticase soy broth Pasteur pipettes Oxalated plasma (preferably rabbit but human plasma can be substituted) Working reagent is prepared by adding 1 vol/plasma to 4 vol/broth in a tube. Inoculation Emulsify a portion of a colony on the side of the tilted tube of working reagent, straighten the tube until the coagulase reagent covers the inoculum and then mix gently. Incubation Incubate for 4 hours in a water bath at 37˚C and then overnight at room temperature. Test Procedure Check the tube for the presence of a clot at 4 hours. The clot should be a firm gel that will not break up on gentle rocking. A clot at this time is a positive test. Negatives at 4 hours should be reincubated for 18-24 hours before being reported as negative. Negative at 4 hours but positive at 24 hours is a positive test. Any test that has not been checked at 4 hours and is negative at 24 hours should be repeated as some strains produce a staphylokinase and are able to lyse their own clot on prolonged incubation. A positive result identifies the organism as Staphylococcus aureus. Animal strains (S. intermedius— dogs and S. hyicus-pigs) can also give positive tube coagulase tests but are rarely a problem with organisms isolated from humans. A negative tube coagulase excludes S. aureus, but does not identify the isolate as S. epidermidis: such isolates are referred to as coagulase negative staphylococci unless further identified. Interpretation Controls Positive Negative Staphylococcus aureus Staphylococcus epidermidis An additional control of an uninoculated tube should be included to detect organism independent clotting and false positive results. Care should be taken if using citrated plasma as any organism that can utilise the citrate will effectively remove the anticoagulant and induce clotting. This will be a false positive. 26 COMPOSITE MEDIA Composite media is a semisolid tubed medium containing glucose, urea, a pH indicator and an H2S detection system. Results for urease, motility, H2S production and gas from glucose can be achieved from a single tube. When used in conjunction with an ONPG test, the information is sufficient to separate non lactose fermenting faecal isolates from selective media into those that require further identification and those that can be discarded. Reagents and Materials Composite Media Tube Straight Wire Inoculation Use a straight wire to sample a single colony or part of a colony. Inoculate the tube with a single straight vertical stab to the bottom of the tube. Incubation Incubate overnight at 37˚C. Urease positive reactions may be detected much earlier. Test Procedure When examining the tubes look for colour, pattern of growth around stab line and the presence of bubbles within the medium. Interpretation See flowchart overpage Uninoculated tubes are orange. Motility: Non motile organisms grow only on the stab line. Motile organisms grow throughout the tube and produce widespread cloudiness. If in doubt—interpret as negative. Urease: Urease positive organisms produce a pink colour. This may look almost purple and be hard to detect if the organism is also H2S positive. H2S: H2S positive organism produce a black precipitate in the medium. For strongly positive organisms this fills the entire tube, but may only be visible around the initial inoculum with some organisms. Gas from Glucose: This can be detected by the presence of gas bubbles within the medium. Care must be taken not to introduce bubbles during inoculation. If in doubt—interpret as negative. Controls Controls for each batch of media should include an H2S positive Salmonella sp, a Shigella sp and an E. coli.. It is prudent to set up a non motile organism (eg Shigella or Klebsiella) as a control whenever a test is being performed as this is a critical test that can be difficult to read. 27 Application A useful flow chart (Fig 1) that utilises the results from the composite media and ONPG is given below. Figure 1 - Flow chart for faecal pathogen screen UREASE pos NOT Salmonella or Shigella neg MOTILITY non motile GAS neg motile IDENTIFY pos NOT Shigella INDOLE pos NOT Salmonella neg ONPG pos H2S pos neg IDENTIFY NOT Salmonella neg IDENTIFY 28 DECARBOXYLASE TESTS ARGININE-LYSINE-ORNITHINE A test to measure the ability of an organism to decarboxylate an amino acid to form an amine under acidic and anaerobic conditions and thus cause a change in pH to alkaline. The decarboxylase tests are used primarily to determine bacterial groups among the Enterobacteriaceae and Vibrionaceae. Reagents and Materials Arginine, Lysine, Ornithine and Control decarboxylase tubes Sterile paraffin Inoculation Transfer a portion of the colony to be tested to the appropriate media and cover with sterile paraffin oil. A control tube without any amino acid should always be tested in parallel with each battery of amino acids to be tested. Appropriate temperature for organisms to grow. ie. Enterobacteriaceae 37˚C, Vibrionaceae 25˚C for up to 4 days. Examine daily. Incubation Interpretation Initially the test changes from deep purple to yellow as a result of acid production from glucose fermentation, and then back to purple if amino acid decarboxylation occurs. The control stays yellow since it lacks an amino acid. Uninoculated media : Control : Positive : Negative : deep purple yellow (indicates acid production from glucose) deep purple (indicates decarboxylation of the amino acid) yellow (amino acid is not decarboxylated). Note: a purple control invalidates the test. Controls Control Organisms Proteus vulgaris Klebsiella Morganella morganii Salmonella typhi Enterobacter cloacae Enterobacter aerogenes Salmonella typhimurium 1 May be variable at times Arginine + + + Lysine +1 + + + Ornithine + + + + 29 DNASE A test to determine the ability of an organism to produce an enzyme capable of degrading DNA. Reagents and Materials DNA agar plate Hydrochloric acid 1M or Toluidine Blue 0.01% Inoculate the culture to be tested heavily over a 1 cm2 area. Several cultures can be tested on the one plate providing they are sufficiently well spaced to avoid false positives. Inoculation Incubation Overnight at 37˚C or 30˚C for up to 2 days. Test Procedure Flood the plate with hydrochloric acid or toluidine blue, leave for 1 minute and then carefully pour off. Interpretation Intact DNA precipitates with strong acid, therefore positives are indicated by a zone of clearing under and around the colony. Negatives are opaque near the colony. Toluidine blue changes to a pink colour with breakdown products of DNA hydrolysis and therefore positive colonies will have a pink halo. Controls Positive Negative Serratia marcescens or Staphylococcus aureus Escherichia colior Staphylococcus epidermidis 30 GELATIN HYDROLYSIS A test to determine the ability of an organism to produce proteolytic enzymes which liquefy gelatin. Used to assist in identification of some Enterobacteriaceae and differentiate between Staphylococcus and Micrococcus.. Reagents and Materials Mercuric chloride solution (plate test only) Gelatin tube media or Gelatin agar plate Straight wire Method One—Tube Test Inoculation Using a straight wire stab the culture to be tested to the bottom of the tube. 24 hours to 14 days at 22-25˚C or 37˚C. An uninoculated control should be incubated simultaneously with the test organism. Incubation Test Procedure Place both tubes (test organism and uninoculated control) in a refrigerator to determine whether digestion of gelatin has occurred. Make the transfer from incubator to refrigerator without shaking the tube. Interpretation A positive test is indicated by a liquid medium after cooling. Method Two—Plate Test Inoculation Spot inoculate a small portion of the colony to be tested on the gelatin agar plate. Up to 4 different organisms can be simultaneously tested on the same plate providing the inoculations are sufficiently spaced to avoid false positives. Incubation 48 hours at 22-25˚C or 37˚C. Test Procedure Interpretation Flood the agar plate with mercuric chloride. Mercuric chloride precipitates any remaining gelatin in the agar thus positives are indicated by a zone of clearing around the colony. 31 Controls Positive Negative Aeromonas hydrophila Klebsiella pneumoniae GERM TUBE Candida albicans is the most frequently isolated yeast from clinical specimens. It can be quickly and reliably identified with the germ tube test. Reagents and Materials Plasma or serum (source not important) Glass culture tube Inoculation Incubation Inoculate a very small amount of yeast into 0.5 ml of serum or plasma. 37˚C for NO LONGER than 3 hours. A water bath is more preferable for incubation. Test Procedure Remove a drop of the suspension, place on a microscope slide, cover slip and examine under the x40 objective for the presence of germ tubes. Interpretation A germ tube is defined as an appendage that is approximately 1/2 the width and 3-4 times the length of the yeast cell from which it arises and does not have a point of constriction at the connection (see Fig 1). For most situations, a germ tube positive yeast can be identified as Candida albicans, however Candida dubliniensis is also germ tube positive and is similar to Candida albicans in other morphological characteristics. Candida dubliniensis is a rtelatively rare isolate and has so far only been associated with disease in HIV patient. Fig 1 Germ Tube budding only positive germ tube Controls Positive Negative Candida albicans Candida tropicalis 32 HIPPURATE HYDROLYSIS Introduction A test to detect the ability of organisms to hydrolyse hippurate (benzyl glycine) to benzoate and glycine. The hydrolysis is detected by adding ninhydrin which forms a coloured end-product with amino acids. The test is usually used in the presumptive identification of Streptococcus agalactiae or in the speciation of Campylobacter. Reagents and Materials 1% Na Hippurate broth 3.5% Ninhydrin solution Inoculation Incubation Transfer a portion of the colony to be tested into the hippurate broth. With a reasonably heavy inoculum, 2 hours at 37˚C in a water bath is sufficient. Alternatively, overnight at 37˚C can be used. Test Procedure Interpretation Following incubation, add 2-3 drops of ninhydrin and wait for 10 minutes. A deep purple colour is positive and no change is negative. For presumptive identification of streptococci, use only on ß-haemolytic colonies and use in conjunction with other tests. A hippurate positive ß-haemolytic streptococcus that gives marginal growth on MacConkey No 1 agar is presumptively identified as Streptococcus agalactiae. It is important to note that both Listeria monocytogenes and Enterococcus faecalis may give positive results. Controls Positive Negative Streptococcus agalactiae OR Campylobacter jejuni Streptococcus pyogenes OR Campylobacter coli 33 HYDROLYSIS OF COMPLEX SUBSTRATES CHITIN, CASEIN, TYROSINE AND XANTHINE A test to determine the ability of an organism to digest complex carbohydrates and/or amino acids. Chitin hydrolysis is a useful test for differentiation of marine organisms, while hydrolysis of casein, tyrosine and/or xanthine are useful for differentiation of aerobic actinomycetes, saprophytic or proteolytic organisms. Reagents and Materials Inoculation Agar plate containing appropriate substrate. Spot inoculate a small portion of the colony to be tested on the appropriate agar plate. Up to 6 different organisms can be simultaneously tested on the same plate providing the inoculations are sufficiently spaced to avoid false positives. Incubation At the appropriate temperature for the organism for up to 30 days. Note: for extended incubations seal the plates with cellotape to avoid excessive moisture loss. Interpretation Hydrolysis of the substrate is indicated by a zone of clearing of the substrate under and around the colony. Brown colouration on the tyrosine plate may also be present for some organisms and is considered positive. Controls Positive Negative Casein Streptomyces griseus Norcardia asteroides Chitin Vibrio parahaemolyticus Escherichia coli Tyrosine Streptomyces griseus Norcardia asteroides Xanthine Streptomyces griseus Norcardia asteroides 34 INDOLE A test to determine the ability of an organism to produce indole from the amino acid tryptophan. Commonly used to assist in the differentiation of Edwardsiella from Salmonella and Escherichia coli from Klebsiella-Enterobacter. Indole broth Kovacs Indole reagent for tube test only Spot Indole reagent (Kovacs without organic solvent) for spot test only. Filter paper (spot test only) Glass rod or wooden tooth pick (spot test only) Reagents and Materials Method One—Broth Test Inoculation Incubation Transfer a portion of the colony to be tested into broth containing tryptophan. Appropriate temperature for the organism to be tested for up to 3 days. (37˚C for 24 hours for most organisms) Test Procedure Transfer a small portion of the broth to be tested into a clean glass tube. Add 2-3 drops of Kovacs indole reagent and mix gently. Negative tests can be reincubated. Do not use spot indole reagent. Interpretation Immediate production of a deep red colour in the top layer indicates indole. A pale orange colour in the bottom layer is negative. Method Two—Spot Test Test Procedure The spot test is only suitable for colonies taken from blood agar or other media known to contain sufficient tryptophan. Place a small piece of filter paper in a petri dish or on a glass slide and add a few drops of spot indole reagent. Using a glass rod or wooden tooth pick transfer a portion of the colony to be tested to the moistened filter paper. Interpretation Production of a brown-red colour on the filter paper within 2-3 minutes is indicative of indole. A positive control should always be used for this test. False negatives may occur with this test. Controls Positive Negative Escherichia coli Klebsiella pneumoniae. 35 METHYL RED A qualitative test for the ability of an organism to produce acid from glucose and overcome the buffering capacity of the media. Some organisms produce more acid than others. To assist in the identification of Escherichia coli and Yersinia spp. This test should always be performed in conjunction with the VP test and utilises the same test media. Reagents and Materials MR/VP Broth Methyl Red pH indicator Inoculation Incubation Transfer a portion of the colony to be tested into MR/VP broth. For up to 5 days. Test Procedure Transfer a small portion of the broth to be tested into a clean glass tube. Add 2-3 drops of Methyl Red and mix gently. Negative tests can be reincubated. Distinct red colour is indicative of copious acid production and is positive. A yellow or orange colour is negative. Interpretation Controls Positive Negative Escherichia coli Klebsiella spp. 36 MOTILITY A test to determine if an organism is motile or non-motile. Some organisms are motile only at specific temperatures and thus this test must be performed under strict conditions. Reagents and Materials Vaseline, Bluetac or plasticine (hanging drop only) Wooden tooth pick (hanging drop only) Coverslip (hanging drop or wet slide mount) Motility media (tube test only) Straight wire (tube test only) Method One—Wet Slide Mount Test Procedure Place a small drop of the bacterial suspension to be tested onto a clean glass slide and mount with a cover slip. View under 400x magnification. To increase contrast rack the condenser down and close the iris diaphragm. Alternatively use phase contrast or dark field microscopy. Interpretation Motile cells should be observed moving against the flow of the broth. Method Two—Hanging Drop Test Procedure Place a small drop of the bacterial suspension to be tested onto a clean glass coverslip. On a microscope slide make a slight ridge of vaseline, Bluetac or plasticine to support the coverslip. Invert the slide over the coverslip, allow the glass to adhere to the vaseline and quickly turn over the slide so the coverslip is uppermost. The drop should then be ‘hanging’ from the centre of the coverslip (Diagram 3). 1 2 3 View under 400x magnification. To increase contrast rack the condenser down and close the iris diaphragm. Alternatively use phase contrast or dark field microscopy. Interpretation Motile cells should be observed moving against the flow of the broth. 37 Method Three—Tube Inoculation Using a straight wire stab the centre of the medium to a depth of approximately 3 cm. At an appropriate temperature for the organism to be tested. ie. 22-25˚C for Vibrio spp. Yersinia spp. and Corynebacterium spp. 37˚C for Enterobacteriaceae and other organisms. An uninoculated control should be incubated. Incubation Interpretation Motile organisms migrate from the stab line and diffuse into the medium causing turbidity. They may exhibit fuzzy streaks of growth. Non-motile organisms accentuate along the stab line; the surrounding medium remains clear. Controls Uninoculated control will remain colourless and clear. 38 NEISSERIA CARBOHYDRATE FERMENTATION A battery of fermentation reactions for the carbohydrates, glucose, maltose, lactose and sucrose is the traditional approach for the speciation of Neisseria sp. Because of their fastidious nature, the fermentation reactions are carried out on cystine enriched trypticase agar slants (CTA). Reagents and Materials A set of 4 carbohydrate agar slants; Glucose—green lid Lactose—red lid Maltose—blue/white lid Sucrose—blue lid Inoculation These tests are usually performed directly from primary selective plates. In such cases, make a heavy suspension of the organism in 0.5 ml saline and add equal volumes to the surface of the slope in each tube. If using a pure culture, a heavy inoculum can be transferred directly to the slope. Incubation Incubate the tubes at 37˚C for 24 hours in a NON CO2 atmosphere. Reactions will often be detectable in 4 hours and will rarely require incubation beyond 24 hours. Interpretation A positive reaction is detected by a colour change from orange to yellow. Neisseria sp tend to produce a colour change only under the first few millimetres of the slope. A completely yellow tube is indicative of contamination and should be viewed with suspicion. No Neisseria sp are able to ferment all four carbohydrates; such a pattern of reaction is suggestive of false positives—most usually due to incorrect incubation in CO2. Results are interpreted as indicated below—the list is not exhaustive but limited to those clinically significant species that may be confused with Neisseria gonorrhoeae. These tests should never be used in isolation, but preferably in conjunction with additional tests. Organism N. gonorrhoeae N. meningitidis N. lactamica N. cinerea M. catarrhalis Growth on selective media§ + + + +* +* Glucose + + + -† Maltose +# + Lactose + Sucrose - §ability to grow on selective gonococcal media *occasionally grow on primary isolation but not on subculture. #occasional maltose negative strains occur. †occasional glucose positive strains occur. Controls Ideally each fresh batch of media should be tested against at least N. gonorrhoea, N. lactamica and N. sicca to ensure correct carbohydrate addition. Controls are not routinely performed for each subsequent use. 39 NITRATE REDUCTION A test to determine the ability of an organism to reduce nitrate to nitrites or free nitrogen gas. Commonly used for differentiation of Haemophilus spp., Branhamella and Neisseria spp. and to assist in identification of Enterobacteriaceae. Reagents and Materials Nitrate broth Reagent A (α-naphthylamine 0.5%) Reagent B (sulphanilic acid 0.8%) Zinc Powder Glass tube or white tile Nitrate strips (Cook’s Method) Method One—The Spot Test. Test Procedure Phase One Place one drop of each reagent onto a white tile. Using a wooden tooth pick or a clean glass rod (do not use a metal loop) transfer a large colony to be tested to the tile and mix well. Interpretation Phase One Formation of a red colour within 1-2 minutes indicates the presence of nitrites and therefore the organism is positive. If the test is negative proceed to phase two. Test Procedure Phase Two Some organisms can reduce nitrites further to a variety of end products which do not react with the indicator reagent and therefore false negatives in phase one must be differentiated from true negatives. Zinc powder will chemically reduce nitrate (if present) to nitrite which will be detected with the reagents which have already been added to the test. Using a wooden tooth pick or a clean glass rod transfer a tiny quantity of zinc powder to the tile and mix well. Interpretation Phase Two Formation of a red colour within 1-2 minutes indicates the presence of nitrates and therefore the organism is negative. No red colour indicates that all nitrates and nitrites have been reduced to unreactive end products and the organism is positive. 40 Method Two—The Broth Test. Inoculation Transfer a small portion of the colony to be tested to nitrate containing media. Incubation 24 hours at an appropriate temperature for the organism. Test Procedure Phase One Observe for gas in the inverted Durham tube. If gas is present and the organism is a fermenter (check OF or TSI), or if no gas is present proceed as follows; Transfer 4-5 drops of the culture to be tested to a clean glass tube. Add 1 drop each of reagent A and B. Interpretation Phase One If gas is present and the organism is a nonfermenter the organism is positive for nitrate reduction. Formation of a red colour within 1-2 minutes after adding Reagents and Materials indicates the presence of nitrites and therefore the organism is positive. If the test is negative proceed to phase two. Test Procedure Phase Two Some organisms can reduce nitrites further to a variety of end products which do not react with the Reagents and Materials and therefore false negatives in phase one must be differentiated from true negatives. Zinc powder will chemically reduce nitrate (if present) to nitrite which will be detected with the Reagents and Materials. Using a wooden tooth pick or a clean glass rod transfer a tiny quantity of zinc powder to the tube and mix well. Note: excess zinc will mask any reaction. Interpretation Phase Two Formation of a red colour within 1-2 minutes indicates the presence of nitrates and therefore the organism is negative. Method Three—Cook’s Method. Inoculation Place a nitrate strip onto a blood agar plate. Using a straight wire stab-inoculate the culture to be tested into the agar approximately 1-2 mm from the edge of the strip. Inoculate a positive control organism on the opposite side of the strip. Incubation 24-48 hours. Interpretation Controls Positive nitrate reduction is indicated by a dark green or brown colour in the vicinity of the stab culture. Positive Negative Escherichia coli Bacillus subtilis 41 NOVOBIOCIN SENSITIVITY Sensitivity to novobiocin is a presumptive test for the identification of Staphylococcus saprophyticus— a common urinary tract pathogen. Reagents and Materials Novobiocin disc (5 µg) Sensitest or Mueller-Hinton agar Inoculation The disc can be included with the therapeutic discs in a standard antimicrobial susceptibility test. Alternatively, a lawn plate (see method) can be prepared on either Sensitest or Mueller-Hinton agar, using an inoculum of approximately 0.5 McFarland. The disc should be placed in the centre of the plate. Incubation In both cases, plates are incubated overnight, in air @ 37˚C. Test Procedure Interpretation Measure the diameter of the zone of inhibition. Interpretation is based on the assumption that the organism is a slide coagulase negative Staphylococcus species isolated from urine. Zone ≤ 16 mm is resistant: presumptive Staphylococcus saprophyticus Zone > 16 mm is sensitive: organism is NOT Staphylococcus saprophyticus Controls Resistant Sensitive Staphylococcus saprophyticus Staphylococcus epidermidis 42 O/129 SENSITIVITY Sensitivity to the vibriostatic agent O/129 is used to distinguish members of the genus Vibrio from other oxidase positive gram negative bacilli, particularly Aeromonas sp and Plesiomonas sp. Reagents and Materials O/129 discs (150 µg) Nutrient, Sensitest or Mueller Hinton agar (may require supplementation 1% NaCl) Inoculation Prepare a lawn plate (see method) using a 0.5 McFarland standard; Nutrient, Sensitest or MuellerHinton agar can be used but it may be necessary to supplement the media with 1% NaCl if the particular Vibrio sp has an absolute requirement for salt. Place the disc in the centre of the plate. Incubation Incubate overnight in air @ 37˚C. It is not necessary to measure the zone of inhibition; any zone of inhibition should be considered sensitive. This interpretation is based on the assumption that the organism is an oxidase# positive Gram negative bacillus. #Vibrio metchnikovii is the only Vibrio sp that is oxidase negative. O/129 (150 µg) sensitive: probable Vibrio sp or Plesiomonas sp. O/129 (150 µg) resistant: organism is NOT a Vibrio sp. Note: A 10 µg O/129 disc may also be tested—sensitivity or resistance to O/129 at this strength is used to assist in speciation of Vibrio sp. Test Procedure Interpretation Controls Sensitive Resistant Vibrio parahaemolyticus Aeromonas hydrophila ONPG (ß-GALACTOSIDASE) A test to demonstrate the presence of the enzyme ß-galactosidase by utilising the compound onitrophenyl-β-D-galactopyranoside (ONPG) which when hydrolysed produces a yellow compound. An alternative substrate p-nitrophenyl-β-D-galactoside may by used. Used to differentiate between lactose delayed from lactose negative organisms. Reagents and Materials ONPG Broth Inoculation Incubation Lightly inoculate ONPG broth with the organism to be tested. 18 to 24 hours. Interpretation Controls A positive test is indicated by an intense yellow colouration of the broth. Negatives are colourless. Positive Negative Escherichia coli Proteus spp. 43 OPTOCHIN SENSITIVITY Sensitivity to optochin is a presumptive test for the identification of Streptococcus pneumoniae. The test can also be used to screen for the presence of the organisms on primary isolation plates. Reagents and Materials Optochin discs Blood agar or blood sensitest agar. Inoculation and incubation The disc can be included with the therapeutic discs in a standard antimicrobial susceptibility test. In this case the plates should be incubated overnight in air @ 37˚C. Alternatively, the disc may be added to the blood plates used for primary isolation from clinical specimens, or on a simple subculture streak plate. The disc should be placed on the plate at the junction of the first and second streak lines (see fig 1). In this case, incubation should be overnight in air, CO2 or anaerobically @ 37˚C. Fig 1 Optochin Plate = optochin disc Test Procedure Interpretation Measure the diameter of the zone of inhibition. Interpretation is based on the assumption that the organism has been confirmed as an α-haemolytic Streptococcus spp.. Zone size ≥ 14 mm: presumptive Streptococcus pneumoniae Zone size 6—14 mm: equivocal result, suggest alternative test or repeat. Zone size ≤ 6 mm: organism NOT Streptococcus pneumoniae Controls Sensitive Resistant Streptococcus pneumoniae Viridans Streptococcus sp 44 OXIDASE A test used for the detection of the oxidase enzymes. Used to assist in the differentiation of Moraxella and Neisseria from Acinetobacter, and Aeromonas, Vibrio and Plesiomonas from Enterobacteriaceae. Tetramethyl-p-phenylenediamine dihydrochloride 1% solution freshly prepared. The reagent should be colourless and should not be used if a deep blue colour is present. Filter paper Glass rod or wooden tooth pick Petri dish lid or base Reagents and Materials Test Procedure Place a small piece of filter paper in a petri dish and soak with freshly prepared oxidase reagent. Some filter papers give a blue colour and these should not be used. Using a clean glass rod or wooden toothpick transfer a portion of the colony to be tested onto the soaked filter paper. Dirty glass or nichrome wire should not be used. Positive and negative controls should always be used. Interpretation Formation of an intense blue colour within 10 seconds is indicative of positive result. Old cultures and those from glucose, nitrate or tellurite containing media, or from selective and/or differential media should not be used. Controls Positive Negative Pseudomonas aeruginosa or Aeromonas sp Escherichia coli 45 OXIDATION—FERMENTATION (O-F) TEST Microorganisms can produce acid from the metabolism of carbohydrates by either or both of two pathways. Fermentation is an anaerobic process whilst oxidation is essentially a strictly aerobic process. This test usually uses glucose as a carbohydrate—although others can be substituted—and determines whether the organism is fermentative or non-fermentative and/or oxidative non oxidative. The test is useful in many situations as a primary taxonomic test—especially with respect to the aerobic Gram negative bacilli. Reagents and Materials Straight wire Sterile paraffin oil One Tube Method 1 tube of media# # = Hugh & Leifson medium (green) Two Tube Method 2 tubes of media# Gram Pos Method 2 x tubes of media* * = Baird-Parker medium (purple) Inoculation O/F tubes should be boiled to drive off residual oxygen and then cooled in ice immediately prior to use. Use a straight wire to stab inoculate the medium to the bottom of the tube/tubes. In two tube tests overlay the medium in one of the tubes with about 1 cm of sterile paraffin oil to maintain anaerobic conditions. Incubation is at 37˚C in air, 48 hours is usually sufficient although in some cases the tubes may need to be examined daily for up to 14 days. Incubation Interpretation Examine the tubes for the presence and position of acid production in both tubes. With both media, acid production produces a yellow colour. One Tube Method acid (yellow) throughout : fermentative acid (yellow) in top 1/4—1/3 : non fermentative but oxidative no acid (yellow) : non fermentative and non oxidative open and paraffin tube both acid (yellow) : fermentative only open tube acid (yellow) : non fermentative but oxidative no acid (yellow)in either tube : non fermentative and non oxidative Two Tube Method Controls Fermentative: Escherichia coli Non fermentative but oxidative: Acinetobacter calcoaceticus subsp. anitratus Non fermentative and non oxidative: Alcaligenes sp 46 PYR TEST This test detects the enzyme L-pyrrolidonyl-peptidase via the conversion of the test substrate, Lpyrrolidonyl-ß-naphthylamide (PYR), into the free amide which produces a red end product when combined with an indicator. Many gram positive cocci are PYR positive, yet the tests remains very useful because it rapidly identifies and differentiates common and important organisms that were previously difficult to separate. Reagents and Materials Substrate: paper discs impregnated with PYR (250 µL of 0.05%) Buffer: 0.05 M (pH 8) Sorensen’s Buffer Indicator: 1% acidified p-dimethyl-aminocinnamaldehyde Inoculation Make a suspension of the organism in 250 µL of buffer to yield approximately a 1.0 McFarland standard or higher. Add a substrate disc and gently agitate. This test relies on pre-formed enzymes, so a heavy inoculum is critical. Incubation Incubate at 37˚C for 30 minutes. Test Procedure Interpretation Add two drops of indicator to the tube after incubation. Orange-red colour development within 5 minutes is a positive test. PYR pos Streptococcus pyogenes Enterococci Nutritionally variant streptococci Staphylococcus intermedius (tube coag pos) Staphylococcus schleiferi (slide coag pos) Staphylcocccus lugdanensis (slide coag pos) PYR neg Other ß-haemolytic streptococci Non enterococcal Group D streptococci Other viridans streptococci Staphylococcus aureus Controls Positive Negative Any of the positives listed above Any of the negatives listed above SALT TOLERANCE A test to determine the ability of an organism to grow in an inhibitory medium containing NaCl. This test is commonly used for characterisation of catalase negative Gram-positive cocci at a NaCl concentration of 6.5%. Other salt concentrations are useful for differentiation of Aeromonas, Plesiomonas and nonfermenting Gram-negative bacteria. Reagents and Materials Inoculation Incubation Broth containing appropriate amount of NaCl Lightly inoculate the broth with the organism to be tested. At an appropriate temperature for the organism to be tested, for up to 5 days. Interpretation A positive test is indicated by turbidity of the broth due to growth of the organism, or if an indicator is used (bromocresol purple) yellow colouration. Occasionally some organisms will be positive but will not produce any colour change. Controls (6.5% NaCl) Positive Enterococcus spp. 47 Negative Streptococcus spp. SPS SENSITIVITY Sensitivity to sodium polyethanol sulphate (SPS) is used to presumptively identify Peptostreptococcus anaerobius—an anaerobic Gram positive coccus. Reagents and Materials SPS discs (1ug) Blood agar Inoculation A standard streak plate on blood agar (or enriched blood based medium) is made and the disc placed in the second streak zone (see OPTOCHIN TEST). Alternatively a lawn plate (see method) can be prepared on similar media using a 0.5 McFarland suspension and the disc placed in the centre. Incubation Incubate overnight @ 37˚C under anaerobic conditions. Test Procedure Interpretation Measure the diameter of the zone of inhibition. The interpretation is based on the assumption that the organism is a strict anaerobe—this can be demonstrated by no growth on an aerobic subculture or sensitivity to metronidazole. It may be convenient to do this by simply adding a 5 µg metronidazole (MTZ) disc to the SPS SENSITIVITY plate. In addition the organism should be a large Gram positive coccus, appearing in chains and having a sweetish but foul odour. Zone≥ 12 mm is sensitive: presumptive Peptostreptococcus anaerobius Zone < 12 mm is resistant: organism is NOT Peptostreptococcus anaerobius Controls Positive Negative Peptostreptococcus anaerobius Peptostreptococcus magnus 48 SUPEROXOL This test is essentially a modified catalase test that uses 30% hydrogen peroxide instead of the usual 3% solution. The test is used mainly for screening selective media for the presence of Neisseria gonorrhoeae, or once isolated, for supporting the identification. Reagents and Materials 30 % hydrogen peroxide Glass tube Capillary tube or wooden tooth pick Test Procedure Use a loop, capillary tube or toothpick to add a portion of a colony to a glass tube containing reagent. Rapid, vigorous and immediate bubbling is a positive test. Weak or delayed bubbling should be interpreted as negative. Interpretation All interpretation is based on the assumption that the organism is an oxidase positive Gram negative diplococcus. A negative test excludes Neisseria gonorrhoeae. A positive test is highly suggestive of Neisseria gonorrhoeae, although very occasional strains of Neisseria meningitidis and Neisseria lactamica—that have been isolated on selective plates—have given positive results. Caution This test relies on the organism being a Gram negative coccus. Many of the catalase positive Gram positive cocci (eg staphylococci) will give strong positive superoxol tests. Controls Positive Negative Neisseria gonorrhoeae Neisseria lactamica 49 TRIPLE SUGAR IRON Triple sugar iron (TSI) medium determines the ability of an organism to produce acid, with or without gas, from the fermentation of glucose, lactose or sucrose. The medium also contains a system to detect the production of H2S. TSI is generally used in the initial characterisation of gram negative bacilli—particularly into broad groups of fermenters and non-fermenters and in the further characterisation of the Enterobacteriaceae. It can also be used for the detection of H2S production in Erysipelothrix rhusiopathiae. Reagents and Materials TSI agar slope straight wire Inoculation A well isolated colony is selected on the primary culture plate and touched with a straight wire. Stab once, vertically deep into the tube—to within a couple of millimetres of the bottom—when the wire is removed the slant is streaked with a back and forth motion. Incubation Overnight @ 37˚C. Interpretation Uninoculated tubes are orange/red. Acid colour change (A) is indicated by yellow, and alkaline (K) or no change is indicated by red. H2S production is indicated by a black precipitate and gas by the presence of bubbles or disruption of the medium. By convention the tubes are read in two parts; colour change on the slant and colour change in the butt. Alkaline slant & alkaline butt (K/K) No carbohydrate fermentation Alkaline slant & acid/black butt (K/A/H2S) Glucose fermented Lactose and sucrose not fermented H2S produced Alkaline slant & acid butt (K/A) Glucose fermented Lactose and sucrose not fermented Acid slant & acid butt (A/A) Glucose fermented Lactose and/or sucrose fermented 50 UREASE A test to determine the ability of an organism to hydrolyse urea to form two molecules of ammonia. A useful test for differentiation of the Enterobacteriaceae especially Proteus spp. Reagents and Materials Urease agar slope Inoculation Incubation Using a straight wire or loop, heavily streak the surface of the agar slope with the organism to be tested 35˚C for up to 6 days. Examine at 2 and 4 hour intervals, and then daily. Interpretation Controls Formation of a pink/red colour on the slope of the agar indicates hydrolysis. Negative test is no colour change (buff to pale yellow colour). Positive Negative Proteus mirabilis Escherichia coli VP (VOGES-PROSKAUER) A test to determine the ability of an organism to produce acetoin from glucose fermentation. Used to assist in differentiation of Klebsiella spp., Enterobacter spp., Escherichia coli, Staphylococcus spp. and Micrococcus. This test should always be performed in conjunction with the Methyl red test which utilises the same test media. Reagents and Materials MR/VP broth α-naphthol 5% potassium hydroxide 40% Glass tube Inoculation Incubation Transfer a portion of the colony to be tested into MR/VP broth. 24 to 48 hours; may require up to 10 days. Some organisms are variable at 37˚C but positive at 25˚C Transfer 10 drops of the culture to be tested into a clean glass tube. Negative culture can be reincubated. Add 5 drops of α-naphthol and mix gently. Add 2 drops of potassium hydroxide and mix gently. Leave the tube standing at room temperature for up to 30 minutes. Note: it is important to add the reagent in the order stated above since potassium hydroxide can form a red colour with peptones in the medium giving a false positive reaction. Test Procedure Interpretation Controls Formation of a pinkish-red colour indicates a positive reaction. Yellow or copper colours are negative. Positive Negative Klebsiella pneumoniae Escherichia coli 51 X AND V FACTOR TESTING Most members of the genus Haemophilus have a requirement for either or both of two specific growth factors; this is usually used to assist in identification. Reagents and Materials X factor discs (haemin) V factor discs (NAD) X+V factor discs X and V factor free test plate (usually a Columbia agar plate, ie a blood agar base plate) Inoculation Prepare a lawn plate using a 0.5 McFarland inoculum. When preparing the inoculum extreme care must be taken not to transfer any free X or V factor present on the primary plate. If using X, V and X+V discs, these should be placed approximately 10 mm in from the edge of the plate and in such a way as to provide maximum distance between the discs (see Fig 1). If using only X and V strips, these should be placed at an angle in the centre of the plate such that the bottom of the strips are approximately 10 mm apart and the tops approximately 30 mm apart (see Fig 2). Incubation Incubate overnight in CO2 at 37˚C. Interpretation Look for the patterns of growth around the specific growth factor discs. Haemophilus species influenzae parainfluenzae haemolyticus parahaemolyticus aphrophilus paraphrophilus ducreyi Factor Requirements X and V V only X and V V only none V only X only Fig. 1 H. parainfluenzae X Fig. 2 H. influenzae V X xv V = growth 52 ANTIMICROBIAL SUSCEPTIBILITY TESTING BY DISC DIFFUSION Background At one level, these tests are simple. Discs containing antibiotics are placed on a lawn plate of an organism and during incubation, the antibiotic diffuses into the agar, inhibits the organism and produces a zone of inhibition. These tests are used to categorise organisms as either “sensitive”—meaning an infection with the organism is likely to respond to treatment with the drug, or “resistant”, meaning that an infection with the organism will probably not respond to treatment with the drug. However, it is important for the student to realise a simple “zone of inhibition” does not necessarily indicate that the organism is “sensitive”, nor does the absence of a zone indicate that the organism is “resistant”. Disc diffusion antimicrobial susceptibility tests must be carried out according to defined methods where the disc strengths, size of inoculum and interpretative criteria for zone sizes have been determined. Two methods are widely used in Australia, they are the National Committee for Clinical Laboratory Standards (NCCLS) method—which was previously called the Kirby-Bauer method, and the Calibrated Dichotomous Sensitivity (CDS) method. A very important point to remember is that the methods are not interchangeable. For example, you cannot use disc strengths recommended for one method and zone sizes from the other. The methods outlined below are abbreviated and as such are only suitable for testing the selected organisms encountered during the practical sessions. For more complete descriptions of the methods students are directed to seek out the appropriate documents. Disc Diffusion by the Calibrated Dichotomous Sensitivity (CDS) Method Media Sensitest agar: Blood sensitest agar: HTM*: for Haemophilus influenzae for rapidly growing non fastidious aerobes for streptococci, enterococci and M. catarrhalis. * Haemophilus test medium with 15 mg/L of fresh haematin and NAD. Inoculum Preparation A straight wire (0.56 mm diameter) is passed vertically once through a typical colony of at least 2 mm in diameter until the wire touches, but does not penetrate, the surface of the agar. The wire is withdrawn and the inoculum is transferred to a 2.5 ml sterile saline. For S pneumoniae the method is modified such that the wire is held vertically and penetrates into an area of confluent growth, this is then moved horizontally in one direction along the edge of the growth for 1 cm. The final inoculum in both situations approximates 10 6-7 cells/ml Plate Inoculation It is preferable to use dry plates, these can be dried with the lid removed in a 37 C incubator for up to 2 hours on the day of the test. The plates are inoculated by flooding with the saline suspension and rocking the plate gently to achieve even distribution. The excess inoculum is removed with a pasteur pipette. 53 The plate is allowed to dry, with the lid off, for up to but not exceeding 45 minutes before the discs are applied. Incubation Conditions The plates are incubated inverted, in AIR at 37˚C for 18-24 hours. The method is modified for S. pneumoniae , Haemophilus influenzae and Moraxella catarrhalis which are incubated in CO2. Interpretation Zone sizes in the CDS method refer to the annular radius ie from the edge of the antibiotic disc to the edge of the zone of inhibition. The general rule for interpretation uses a standard breakpoint of 6 mm. Zones > 6 mm indicate sensitivity. Zones < 6 mm indicate resistance. Exceptions to the 6 mm rule are; 4 mm with the aminoglycosides and Pseudomonas aeruginosa. 4 mm with vancomycin and Staphylococcus sp. 4 mm with enterococci and ampicillin and gentamicin. 2 mm with vancomycin and enterococci. Disc Strengths The following discs are calibrated for use with the CDS method. Enterobacteriaceae Ampicillin 25 Augmentin 60 Cefotaxime 5 Ceftazidime 10 Gentamicin 10 Tetracycline 30 Trimethoprim 5 Sulphafurazole 300 Naladixic acid 30 Imipenem 10 H. influenzae Ampicillin 5 Cefotaxime 0.5 Chloramphenicol 10 Staphylococci Penicillin 0.5 Methicillin 5 Erythromycin 5 Tetracycline 30 Gentamicin 10 Vancomycin 60 Pseudomonas sp Ceftazidime 10 Gentamicin 10 Ciprofloxacin 2.5 Streptococcus pneumoniae Penicillin 0.5 Cefotaxime 0.5 and 5 Chloramphenicol 30 Erythromycin 5 Tetracycline 30 Enterococci Ampicillin 5 Gentamicin 200 Vancomycin 5 References 1) Bell SM. Antibiotic Sensitivity Testing by the CDS method. Clinical microbiology Update Programme 1984; 21. 2) Bell SM. CDS Users Group Newsletters 1-8. The CDS Users Group can be contacted at the Prince of Wales Hospital, High Street, Randwick, NSW 2031. 54 Disc Diffusion by the NCCLS Method Media Mueller-Hinton agar. Mueller-Hinton agar supplemented with 5% sheep blood for fastidious organisms. Haemophilus Test Medium for haemophili. Inoculum A suspension of organisms is made in 2.5 ml saline until the turbidity matches that of a 0.5 McFarland standard. This provides approximately 108 cells/ml. It is preferable to use dry plates, these can be dried with the lid removed in a 37˚C incubator for up to 2 hours on the day of the test. A sterile cotton swab is dipped into the inoculum and then rotated against the side of the tube to remove excess. The plate is inoculated by streaking over the entire surface of the plate. This is repeated by streaking two more times, rotating the plate 60˚ each time. The plates can be dried for no more than 15 minutes before the discs are applied. Plate Inoculation Incubation Conditions Plates are incubated in air at 37˚C. Haemophilus and Streptococcus pneumoniae are incubated in CO2. Interpretation and Disc Strengths Zone diameter is measured and three categories are used—resistant, intermediate and sensitive. Zone Sizes in mm and Interpretation Antimicrobial Ampicillin 10 Penicillin 10 Methicillin 5 Gentamicin 10 Erythromycin 15 Tetracycline 30 Chloramphenicol 30 Trimethoprim 5 Sulphafurazole 300 Cefotaxime 30 Imipenem 10 Ciprofloxacin 5 Ceftazidime 30 Vancomycin 30 Resistant ≤ 13 ≤ 28 ≤9 ≤ 12 ≤ 13 ≤ 14 ≤ 12 ≤ 10 ≤ 12 ≤ 14 ≤ 13 ≤ 15 ≤ 14 ≤9 Intermediate 14—16 10—13 13—14 14—22 15—18 13—17 11—15 13—16 15—22 14 - 15 16 - 20 15—17 10—11 Sensitive ≥ 17 ≥ 29 ≥ 14 ≥ 15 ≥ 23 ≥ 19 ≥ 18 ≥ 16 ≥ 17 ≥ 23 ≥ 16 ≥ 21 ≥ 18 ≥ 12 References 1) National Committee for Clinical Laboratory Standards. Performance standards for antimicrobial disk susceptibility tests. 5th ed. Approved Standard. NCCLS publication M2-A5. Villanova, pa.:NCCLS 1993. 55 ALBERT’S STAIN—VOLUTIN GRANULES This stain is used to demonstrate metachromatic granules in bacteria eg. Corynebacterium diphtheriae. Reagents and Materials Albert’s stain Lugol’s iodine Method 1. 3. 4. 5. 2. Prepare a bacterial film and heat fix. Flood the slide with Albert’s stain. Leave for 5 minutes. Rinse with a gentle stream of water. Flood the slide with Lugol’s iodine. Leave for 1 minute. Wash the slide with a gentle stream of water. Blot dry. Interpretation Metachromatic granules—black Protoplasm and other bacteria—green CAPSULE AND NEGATIVE STAINS These are negative stains in which the background is stained and the capsule appears colourless. Reagents and Materials Indian ink Glucose 6% Crystal violet 57 Method One—Dry India Ink 1. 2. 3. 4. 5. 6. 7. 8. 9. Place a loopful of 6% glucose at one end of the slide. Add a small amount of bacterial culture to the glucose and mix well to form an even suspension of cells. Add a loopful of India ink to the drop and mix. With the edge of another glass slide spread the mixture of ink, glucose and cells over the slide. Allow the film to dry in air without heat. Fix the film by flooding the slide with methanol. Drain off the excess. Dry the slide by gently waving it in the air. Flood the slide with crystal violet. Leave for 30 seconds. Wash the slide with a gentle stream of water. Gently blot the slide dry. Interpretation Organisms—purple Background—dark grey Capsule—clear halo. 58 Method Two—Wet India Ink 1. 2. 3. 4. Place a loopful of India ink on a clean slide. Emulsify a small colony in a drop of distilled water, mix with the drop of ink. Place a clean coverslip on the drop and press down firmly with a piece of blotting paper to give a thin film. Avoid contaminating your fingers and discard the paper into disinfectant. Examine with the 40x objective with a reduced light source. Interpretation The background is dark grey, the organisms are unstained and the capsule stands out as a shiny ring around the organism. FLAGELLA STAIN This procedure can be used to demonstrate the number and arrangement of flagella on bacterial cells. Care must be taken at all times to avoid disrupting the cell and flagella. Solution 1 contains 10 ml of 5% aqueous phenol, 10 ml of saturated aqueous aluminium potassium sulphate-dodecyl hydrate and 2 g of tannic acid. Solution 2 contains saturated ethanolic solution of crystal violet. For staining a mixture of 10 parts of solution 1 is mixed with 1 part of solution 2 and filtered. The final stain is best used from a syringe fitted with a 0.22 µm filter. Reagents and Materials Method 1. 2. Place a drop of sterile water onto a clean glass slide. Using an applicator stick or straight wire, touch the margin of a colony of motile organisms and then transfer the stick or wire into the drop of water on the slide. Do not stir the stick or wire in the drop. Allow the organisms to swim into the water droplet. Do not allow the slide to dry out. Alternatively, using a loopful of water gently touch the colony margin allowing motile cells to swim into the loop. Apply the loopful of water with cell to the slide. 3. 4. 5. 6. Place a clean glass cover slip onto the water droplet and examine for motile cells. Leave the slide for 10 minutes to allow the cells to attach to the slide or cover slip. Using a loop apply 2 drops of stain to the edges of the cover slip. Allow the stain to flow under the cover slip by capillary action. Leave for 10 minutes. Seal the edges of the coverslip with nail polish. Examine the slide with the x100 objective for bacterial cells and flagella. Note: if you wish to delay examination leave the slide in a sealed moist chamber. For phase contrast microscopy dilute the crystal violet solution 1/10 prior to making the final stain. Interpretation Bacteria stain purple and flagella can be seen as faint blue appendages. 59 GRAM STAINING This is the most important differential technique applied to bacteria and with a little care and practice will contribute greatly to your understanding of microbiology. Crystal violet binds to an iodine mordant to form a large complex which is prevented from leaving the Gram-positive cell when decolourisation occurs. Gram-negative organisms contain substantially more lipids in the cell wall, which dissolve allowing the crystal violet-iodine complex to wash away and thus the cell can stain with the counter stain. Reagents and Materials Crystal violet Gram’s Iodine Acetone/alcohol Carbol fuchsin Method 1. 2. 3. 4. 5. 6. 7. 8. 9. Prepare a bacterial film and heat fix. Flood the slide with crystal violet for 30 seconds. Drain and wash the slide with a gentle stream of water for approximately 2 seconds. Rinse and flood the slide with iodine. Leave for 1 minute. Drain and wash the slide with a gentle stream of water . Holding the slide at an angle run alcohol-acetone over the surface until no more colour emerges— about 2 seconds. Take care it is easy to over decolourise. Wash immediately with water; drain off excess. Replace slide on staining rack and flood with counterstain—safranine, dilute carbol fuchsin or neutral red, 30 seconds. Drain the slide and rinse with a gentle stream of water. 10. Carefully blot the stained slide between pieces of blotting paper. Interpretation Gram-positive bacteria and yeasts—blue/purple Gram-negative organisms, tissue cells and protein—red/pink. 60 KINYOUN STAIN This is an acid fast stain used to detect acid fast parasites in faecal specimens. Reagents and Materials Concentrated carbol fuchsin (Kinyoun stain) 3% acid alcohol Counterstain (Malachite green 0.5% or methylene blue 0.3%) Method 1. 2. 3. 4. 5. 6. 7. 8. Prepare a thin smear of faeces—air dry and gently heat fix. Flood smear with carbol fuchsin. Leave for 30 minutes. Apply additional stain as required to prevent the slide drying out. Wash the slide in a gentle stream of water. Briefly decolourise with acid alcohol until no more stain comes off. The time for decolourisation will vary with the thickness of the smear—but it should not be longer than about 3 mins. Wash the slide in a gentle stream of water. Flood the slide with counterstain for 30 seconds. Wash the slide in a gentle stream of water. Gently blot and air dry. Scan the slide using the x 40 objective and confirm acid fast objects using oil immersion. Interpretation Cryptosporidium parvum oocysts are round, about 4-6 um in diameter and stain red (acid fast). Cyclospora cayatenensis oocysts are also acid fast, but are usually 10-14 um in diameter and characteristically show variability in stain retention: some oocysts stain strongly acid fast, some weak and some not at all. Control A positive control slide—known to positive for Cryptosporidium—should be run with each batch. 61 LACTOPHENOL COTTON BLUE TAPE PREPARATION This technique is excellent for examining the microscopic morphology of molds. The advantage of this technique over others is that it retains the fungal architecture that is so important in identification. Reagents and Materials Lactophenol cotton blue Alcohol Forceps Sticky tape Method 1. 2. 3. 4. 5. 6. Dip tip of forceps in alcohol Cut off 1 cm x 1 cm square of sticky tape and hold with forceps. Apply sticky tape to surface of mould—sticky side down. Place sticky tape onto glass slide—sticky side up. Apply 1 drop of alcohol and then 1 drop of stain to the sticky tape. Coverslip. METHYLENE BLUE (SIMPLE) STAIN This stain is easy to perform and can be used to demonstrate cell size, shape and arrangement. Reagents and Materials Loeffler’s methylene blue Method 1. 3. 2. Prepare a bacterial film and heat fix. Flood the slide with methylene blue. Leave for 1 minute. Wash the slide in a gentle stream of water. Blot dry. Interpretation Cells will appear light blue in colour. 62 SPORE STAIN Often it is not necessary to use any special technique for demonstrating spores since they can be seen in the Gram stain as non-staining ovoid or spherical bodies lying within the stained bacteria. They can be seen far more easily using phase contrast microscopy of a wet preparation when they appear as bright refractive granules. They nature of the spore necessitates a vigorous staining procedure but once stained it resists decolourisation. Note: use an old culture for demonstration of spores since some organisms require several days before spores will be produced. Reagents and Materials Malachite green 5% Alcohol for heating Safranine 0.5% Method 1. 2. 3. Prepare a bacterial film and heat fix. Flood the slide with 5% malachite green. Using a heating wand, heat the slide gently from underneath until the steam rises. Allow the slide to stain for 5 minutes. Intermittent heating may be required. Do not allow the slide to dry out. If necessary use more stain. 4. 5. 6. Wash the slide with a gentle stream of water. Flood the slide with 0.5% safranine. Leave for 30 seconds. Wash the slide with a gentle stream of water and gently blot dry. Interpretation Spores—green Protoplasm or vegetative cells—pink/red 63 ZIEHL-NEELSEN (ZN) STAIN The ZN stain is required for staining organisms with a large quantity of lipids (mycolic acids) in their cell wall which prevent other stains from penetrating the cell. Concentrated carbol fuchsin is heated to enable the stain to penetrate the cell and decolourisation utilises an acid/alcohol mixture. Acid fast cells are those which resist decolourisation, thus the stain is fast within the cell. Reagents and Materials Concentrated carbol fuchsin Alcohol for heating Acid/alcohol Malachite green 0.5% or methylene blue 0.3% Method 1. 2. 3. 4. 5. 6. 7. 8. 9. Prepare a bacterial film and heat fix. Ensure the slide rack is even before applying the stain. Flood the slide with concentrated carbol fuchsin. Using a heating wand, heat the slide gently from underneath until the steam rises. Start timing from this point. Allow the slide to stain for 5 minutes. Do not allow the slide to dry out. Wash the slide in a gentle stream of water. Flood the slide with acid/alcohol. Leave for 4 minutes. Thoroughly wash the slide in a gentle stream of water. Flood the slide with 0.5% malachite green or 0.3% methylene blue. Leave for 30 seconds. Wash the slide in a gentle steam of water. Gently blot dry. Clean the underside of the slide of carbon deposit from the heating wand. Interpretation Acid fast cells—red Non acid fast cells—green or blue depending upon which counter stain is used. Note: inadequate washing after decolourisation will cause the counter stain to slightly change colour. Thus malachite green stained features will appear slightly blue. 64