Clinical Research Associates

2. ANTIMICROBIAL TESTING USING ORAL BACTERIA: Desensitizers
OVERVIEW
This test is designed to assess potential antimicrobial ability of various desensitizers using a modification of the Kirby-Bauer test. The Kirby-Bauer test was developed to assess antibiotic resistance of microbes in clinical samples by measuring the zone of inhibition of microbial growth around disks soaked with a known amount of antibiotic.
MATERIALS
Equipment:	Beakers, glass Biohazard Buckets Biological Safety Hood Ethanol Burner Fleakers, 1 liter Forceps, sterile Glass rod, bend (glass hockey stick for spreading) Incubator, 35°C M25 Microman (Gilson) Pipette bulb or autopipetter Ruler, metric Test Tube Racks, 80 place, 20 x 150 mm and 25 x 150 mm Test Tubes, sterile, 20 x 150 mm and 25 x 150 mm Timer Vortex Mixer

Supplies:	Aluminum Foil Autoclave Tape Bags, Biohazard, 12 x 24 inch Bags, Sterilization Capillary Tips & Pistons, CP-25, sterile Ethanol, 70% & 95% Filter paper disks, Grade 740-E Special-Purpose (VWR), 0.64 mm diameter Gloves, nonsterile, power free latex preferably Masks Matches or lighter Petri Dishes, sterile, 100 x 15 mm Pipettes, glass, sterile, 1 ml Towels, paper

Media:	Tryptic Soy Agar Powder (Becton-Dickinson) Tryptic Soy Agar + 5% Sheep’s Blood Agar Plates, (Allegiance, product #432239) Sodium Chloride (NaCl) for Physiological Sterile Saline Water, sterile deionized (DI)

Organisms:	Oral bacteria in saliva collected fresh from multiple volunteers


METHODS
I. Test Preparation:
	A.^°	Place glass hockey stick and filter disks in sterilization bags and place in a steam sterilizer at 125^°C for 30 min.
	B.^°	Place capillary pipet tips and pistons in tip holder boxes and place in a steam sterilizer at 125^°C for 30 min.
II. Media Preparation
	A.	Prepare agar plates
		1.	Typtic Soy Agar + 5% Sheep’s Blood Agar (SBA).
			a.	SBA agar can be made in the lab, or ordered pre-poured from a commercial source.
			b.	Pre-poured agar needs to be refrigerated as soon as it is received until immediately before use.
			c.	CRA uses pre-poured agar plates the same week they are ordered
		2.	Tryptic Soy Agar (TSA):
			a.	Add 20 g Tryptic Soy Agar to 500 ml DI water in a 1-liter fleaker.
			b.	Mix with heating until solution begins to boil. Solution will go from opaque to clear right before boiling begins.
			c.	Autoclave for 30 min on a liquid cycle.
			d.	Place fleakers on stir plates. Cool with continuous stirring.
			e.	When cooled to approximately body temperature (fleakers feel slightly warm or “baby bottle” warm), aseptically pour about 23 ml into 100 x 15 mm Petri dishes.
			f.	Plates need to be used within 30 - 60 min of pouring.
	B.	Prepare Physiological Sterile Saline (PSS).
		1.	Add 8.5 g NaCl to 1000 ml of deionized water.
		2.	Aliquot into 500 ml plastic bottles.
		3.	Autoclave 15 – 30 min at 121°C.
III. Saliva Collection
	A.	Select volunteers.
		1.	At least four volunteers are needed. More is better.
		2.	Volunteers should not have used any antimicrobial substance before saliva collection.
			a.	No Listerine, antibiotics, etc.
		3.	It is preferable, but not necessary, that the volunteers have not recently eaten.
	B.	Collect saliva.
		1.	Give sterile, capped culture tubes to volunteers, one per volunteer.
		2.	Instruct volunteers on how to donate.
			a.	Only saliva is needed. Instruct the volunteers not to donate mucus. Mucus would contain upper respiratory bacteria, not normal oral flora.
			b.	The greatest amount of non-foamy salvia seems to be generated by ‘milking’ the sublingual saliva glands to form a pool under the tongue, then gently spitting the saliva into the tube.
			c.	Gum chewing to stimulate saliva productions was not a cause for discarding the saliva, but is discouraged.
			d.	Volunteers should donate 1 – 5 ml, depending on the number of desensitizers to be tested.
	C.	Store the saliva in a refrigerator until testing.
IV. Testing
	A.	Label the plates.
		1.	Three TSA and three SBA plates are needed for each desensitizer.
		2.	On the bottom of each plate, write the following:
			a.	Test date
			b.	Desensitizer name
			c.	Replication letter (A, B, or C)
			d.	Since TSA and SBA are easily distinguished by color, the media designation is not necessary
	B.	Inoculate the plates.
		1.	Wipe down the inside surfaces of the biosafety hood with 70% ethanol.
		2.	Wipe down with 70% ethanol the equipment & containers of supplies needed in the hood:
			a.	Vortex
			b.	Biohazard bin with biohazard bag
			c.	Microman and capillary tip box
			d.	Ethanol burner
			e.	Beaker
			f.	Bottle of 95% ethanol
		3.	Place equipment in the hood.
		4.	Place sterile, bagged equipment and supplies in the hood:
			a.	Glass hockey stick
			b.	Forceps
			c.	Individually wrapped 1 ml glass pipets
			d.	Sterile filter disks
		5.	Combine the saliva samples in the hood by pouring into a sterile container, such as a test tube.
		6.	Place labeled plates in the hood.
		7.	Pour 95% ethanol into beaker.
		8.	Light ethanol burner.
		9.	Aseptically remove hockey stick from sterilization pouches.
		10.	Using a 1 ml glass pipet, place 0.2 ml of saliva on a plate.
			a.	Saliva tends to be too viscous to aliquot using a Pipetman. A glass, 1 ml pipet is accurate and is effective.
			b.	Vortex the saliva before removing aliquots.
		11.	Use the sterile hockey stick to spread the saliva evenly over the surface of the plate.
			a.	This is easiest if the saliva is placed in the center of the plate.
			b.	After spreading, do not turn the plates over. The saliva takes >4 hours to soak into the agar and will run before than.
		12.	Set plate, right side up, in a clear area of the hood where it will not get in the way.
		13.	Repeat steps IV.B.10 – 12 on remaining plates.
		14.	Sterilize glass hockey stick if contaminated by ‘flaming.’
			a.	Dip spreader end of stick into beaker containing 95% ethanol.
			b.	Before ethanol drips off, place stick in open flame.
			c.	Watch to make sure blue flames cover the entire surface to be sterilized.
			d.	Do not hold wet stick over beaker full of ethanol when flaming. Burning drops of ethanol can drop into the beaker and set the contents on fire.
			e.	Allow a few seconds to cool before using to spread saliva.
	C.	Place desensitizer on plates.
		1.	Aseptically open pouch containing filter disks.
		2.	Aseptically open pouch containing forceps.
		3.	Hold one disk in sterile forceps and, using the M25 Microman, aseptically place 10ul of desensitizer on filter.
		4.	Using the forceps, place the desensitizer soaked filter upside down in the center of a saliva inoculated plate.
		5.	Use the forceps points to gently tap the filter down, to insure good contact with the agar.
		6.	Repeat steps IV.C.1 – 5 five more times, so that there are three plates of each kind with disks soaked with that particular desensitizer.
		7.	Flame the forceps before moving onto the next desensitizer (see steps IV.B.14).
		8.	Place plates, right side up, in a clear area of the hood where they will not be in the way.
	D.	Controls.
		1.	Repeat steps IV.C. with the following differences:
			a.	Place a filter disk on a TSA plate and a SBA plate without adding anything to it. This is the 'dry’ control. It is to show that the filter disk will not inhibit the growth of the oral bacteria on the agar.
			b.	Aseptically place 10ul of PSS on disks rather than a desensitizer. This is the PSS control, to show what no zone of inhibition should look like. Perform two PSS controls each for TSA and SBA plates.
	E.	Incubate plates.
		1.	Once all the plates have been inoculated with saliva and have a desensitizer soaked disk placed on it, put the places in a 35°C incubator.
		2.	After ~12 hours, the saliva will have soaked into the agar and the plates can be turned over.
V. Measuring zones of inhibition
	A.	Zones should be measured at ~24, 48, & 72 hours after start of incubation.
	B.	At the above times, remove the plates from the incubator and place in a biosafety hood.
		1.	Put on gloves and mask before handling plates.
		2.	For comfort in working with the plates, be sure the hood vents HEPA filtered air to the outside and not back into the room. The plates will emit an odor like halitosis.
		3.	Put plates in hood upside down.
	C.	Measure and record the zones of inhibition.
		1.	Oral bacteria should have grown to cover the surface of the plate, except, potentially around the disk
		2.	If the desensitizer had prevented the growth of the oral bacteria, there will be a clear space around the disk with no growth. The clear space is called the zone of inhibition.
		3.	Hemolysis.
			a.	There may also be an area on the SBA plates were the red of the blood cells is gone, leaving the tan of the TSA. This is called ‘b-hemolysis.’ The disinfectant has lysed the red blood cells completely.
			b.	If there is an area of green-brown discoloration, this is a-hemolysis, in which the red blood cells have not been completely lysed.
			c.	The zone of inhibition may or may not cover the same area as the hemolysis.
		4.	Using a metric ruler, measure the diameter of the zone of inhibition. This measured most accurately through the diameter of the disk. Because the zone is not always a perfect circle, measure in at least two directions, perpendicular to each other.
		5.	Record the measurements.
		6.	Check the controls.
			a.	Both the dry disk and the disks soaked with PSS should have a ‘0’ zone of inhibition or a lack of a clear space in the bacteria growth.
			b.	If there is a measurable zone of inhibition, this should be subtracted from the test zones of inhibition.
		7.	Replace the plates in the incubator until the next time for measuring the zones of inhibition.
VI. Analyze the Data
	A.	The larger the diameter of the zone of inhibition, the greater the disinfecting potential.
	B.	Analyze the data statically.
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