Links to files on the current site

Home

Syllabus

• Agar slant tubes
• Aseptic technique
• Assays
• Assays Illustrated
• Colony morphology
• Gram stain
• Isolating colonies
• Making serial dilutions
• Culture media
• Preparing spread plates
• Relationship to oxygen
• Viewing Gram stains
• Viewing living bacteria

• Characterization writeup
• Lab responsibilities
• Project proposal guidelines
• Using Bergey's

Supplemental pdfs

• Inoculating sticks
• Microscopy procedures

Preparing culture media

Bacteria display a wide range of nutritional and physical requirements for growth. The following requirements, however, are required by all bacteria.

• A substrate – they have to grow in or on something, such as liquid, on an agar surface, on or in an animal host, etc.
• Water – bacteria can be freeze-dried and remain viable, but without water they won't grow
• A source of energy – usually a carbon source, but some bacteria can utilize inorganic compounds and some are photosynthetic
• Source(s) of carbon, nitrogen, sulfur, phosphorus
• Inorganic ions, e.g., Ca²⁺, Mg²⁺, Na⁺

Most bacteria have additional requirements such as for vitamins and growth factors, and many species are quite fastidious, meaning that they have complex nutritional requirements. We will isolate "cultivable" bacteria from agar plates and store them in tubes containing agar media. Agar is a complex carbohydrate extracted from marine algae that solidifies below temperatures of 45˚C. It is not a nutritional component. Pre-mixed dehydrated agar media contains complex nutrients in addition to 1.5% agar. We will use R2A agar media to isolate and maintain bacteria and will also grow our isolates on tryptic soy agar.

About dehydrated media

Pre-mixed dehydrated media comes in the form of granules or powder. We rehydrate the media acording to instructions on the label, by mixing a measured amount of medium per measured volume of deionized water. These materials are very expensive, typically more than $200 a kilogram, and tend to be very hygroscopic. To prevent our media from ruin in our very humid environment, we keep the bottles tightly closed with a couple of silica gel packets inside to take up moisture.

Sterilizing media

Any surface that is exposed to the air will harbor bacteria, fungi, and their spores. Therefore we use presterilized petri dishes for our agar plates and sterilize all supplies and media used to grow and handle our bacterial cultures. We sterilize most media and supplies using a steam autoclave to produce moist heat. We may employ other methods, including filtration, ethylene oxide, radiation, or ultraviolet light as needed.

An autoclave is designed to deliver steam into a pressure chamber, generating high heat and pressure at the same time. High pressure (typically 20 lbs/sq. in) allows the temperature to exceed 100 degrees without boiling off the water from the solutions being sterilized. Heating media to above 121˚C for a sufficient time destroys all living cells and spores. We use an autoclave that starts timing when the temperature reaches 121 degrees, and exhausts the steam slowly after the prescribed time above 121 degrees (to prevent exploding bottles!). The autoclave is effectively a giant pressure cooker. We determine sterilization time by trial and error. Larger liquid volumes require more time than smaller volumes.

To properly use an autoclave

• Most autoclaves are shared equipment – be a good "lab citizen" by learning and following the local rules
• Learn how to properly (and safely) use the instrument
• Because a cycle can take an hour or more, check with others in the lab before starting a run; they may also have something ready to go in
• Do not turn the instrument off or leave the door open
• Be well aware that all inside surfaces are hot, as is anything coming out of an autoclave; exercise appropriate precautions

Preparing agar plates

Different labs will use different procedures depending on their needs. The steps below describe how to prepare tryptic soy agar (TSA) or Reasoner's 2A agar (R2A agar). Please plan to view the accompanying video when it is made available.

1. Place 18.2 grams of dehydrated R2A agar (or 40 grams of dehydrated TSA) into a clean 2 liter erlenmeyer flask, and add 1 liter of deionized water. As you pour the water in, be sure that it washes down any powder that is clinging to the sides of the flask.
2. Swirl the flask vigorously to mix the powder into the water as homogeneously as possible.  It's not usually possible to make it completely homogeneous; don't worry about a few small lumps.  Cover the flask with a foam stopper or aluminum foil, put it in an autoclave tray, and place immediately into the autoclave. Don't add the water until you are ready to put the flask into the autoclave.
3. Autoclave on a liquid cycle, using the recommended program (we sterilize for at least 30 to 45 min above 121˚C)
4. When the cycle is complete, use autoclave gloves to remove the tray from the autoclave.  Still using autoclave gloves, lift the flask to eye level and check that you have a homogeneous clear solution.  Whether or not it appears completely clear, swirl the flask gently and look to see if areas of inhomogeneity appear.  Continue swirling until you have a clear homogeneous solution.
5. Place the flask into the 50 degree water bath.  Note: When grasping the flask, avoid letting the gloves come in contact with the rim of the flask.  If you contaminate the rim, that contamination can get into the agar as you pour it from the flask.  Allow the flask to cool in the 50 degree bath for 40 to 60 minutes before pouring.
6. Set up the plates that you will need in the laminar flow hood in stacks of 5 plates (fewer if your hands are smaller).  Set up the stacks in a row down the middle of the hood.  When ready to pour, bring each stack, in turn, to the near edge of the solid floor of the hood, where it's easier to reach them.
7. Take the flask out of the 50 degree bath (again being careful not to touch the rim). With paper towels, wipe off the water from the outside of the flask (otherwise that contaminated water can drip into your plates). Wrap a paper towel around the neck of the flask, slightly below the rim, and hold that towel in place as you pour (this is to catch dribbles that roll down the outside of the neck, and keep these, now contaminated, dribbles from falling into the next plate that you pour).
8. Raise the front shield of the hood enough that you can comfortably get the flask in to pour. For each stack, pour the bottom plate first, lifting its cover along with the 4 upper plates. Then replace its cover, and pour the second plate from the bottom in the same way. Repeat for each plate in the stack, and then do the same for the next stack.
9. One liter of agar will make 30 to 40 plates, depending on how liberally you pour them.  Pour gently to avoid making bubbles and to avoid overfilling the plate. You can get just the right amount (~25 ml//plate) by pouring from one side until about a nickel-sized area of the plate remains uncovered. Stop pouring and let the agar close up on its own.
10. When finished, label the top plate in each stack with the type of agar (e.g., R2A), the date, and an ID such as initials or team number. Slide the plates to a back corner, being careful not to tip or bounce them so the liquid splashes out. This is to leave the front space available for use by others. If nobody else is waiting to use that space, you can leave the plates where they are until they have solidified, so you don't have to worry about spilling. Once solidified, stack them in taller stacks, to take up less room, and put them in a back corner. 
11. Allow plates to cool and lose some moisture; best practice is to leave closed in a hood for a day or so, then store plates inverted in a closed container.
12. Aside from contamination from dust particles due to careless handling, condensation and insect contamination are our worst enemies; usually we do not refrigerate plates. Immediately discard any plates that show signs of contamination, including mold or fruit fly larvae.

NOTES

• We use erlenmyer flasks of double the agar volume so that there is a large surface area, allowing heat to penetrate more quickly than it would if we filled the container; the extra air space also provides room for the liquid to expand
• Despite all precautions we sometimes see colonies growing on supposedly sterile plates after a few days; you might incubate a couple of plates from each batch at 30˚C for a couple of days, and if they grow colonies discard the entire batch and report the problem

Preparing media in tubes

The only difference between broth and agar media is that broths do not contain an agar component. We use broth tubes primarily for specific assays, or (rarely) for bacteria that will not form colonies on a solid surface. In broth a species may display characteristics that we do not see in preparations from agar cultures. We store isolates in agar slant tubes, prepared by sterilizing agar in a tube then tilting the tube on its side while it solidifies, in order to create a large slanted surface for growth. Slant tubes usually have a couple of cm of solid agar at the bottom, with the top of the slanted surface stopping a cm or so below the top of the tube. Below are the procedures that we employ in our laboratory.

1. Determine the volume that you need per tube using measured amounts of deionized water, then calculate the amount of medium to prepare (number of tubes X volume/tube + about 10% extra)
2. If you are preparing agar slant tubes then use tubes with distilled water to plan for the best volume and angle for tilting the tubes
3. Place your tubes in an autoclaveable rack
4. To prepare agar for a tube culture mix the agar in a large beaker rather than a flask, drop in a stir bar, and heat to boiling in a microwave oven (watch carefully to prevent boiling over). Stir to distribute the agar uniformly and if necessary heat some more and stir until you have a clarified solution. We usually do not need to heat broth media.
5. Distribute the medium into tubes; we have found a large syringe to be a convenient tool for quickly filling tubes; keep the media stirring while filling tubes, especially when preparing agar medium, to keep it well mixed and homogeneous
6. If you are using plain culture tubes then place a vented closure onto each tube, pushing it all the way down; if you are using screw capped tubes, then onto each tube screw the cap all the way down then back off a half turn to allow venting
7. Sterilize on a liquid cycle for the appropriate time
8. Remove the racks and allow to cool before inoculating the media; if you are preparing slant tubes then let them cool while tipped at the predetermined angle

NOTES

• We use slant tubes to store isolates because the small opening minimizes the risk of introducing airborne contaminants and the small air volume inside helps the agar hold moisture, keeping the culture viable long after a plate culture would have dried out
• Media are expensive – do not prepare far more than you need and throw most of it away or (worse) leave it in a nonsterile container for someone else to clean up after it grows mold

Using a sterile cabinet

Unlike a fume hood, which is designed to keep airborne substances from escaping into the laboratory environment, a sterile cabinet keeps airborne contaminants from getting into the hood. A simple laminar flow hood protects exposed sterile surfaces that are placed inside. A containment hood does both jobs, keeping airborne particulate matter from going in or out. To use a hood properly, remember these points.

• Keep all surfaces clean and dry
• Frequently use the UV light to sterilize the interior surfaces; do not stare at the light, which can cause retinal damage
• The opening must not exceed the recommended sash height
• Surfaces kept to back of the hood are more likely to remain sterile, as are objects kept close to the table surface
• Keep non-sterile objects closer to the front, sterile objects to the back
• Keep the hood fairly uncluttered
• Never reach over a sterile surface - you WILL contaminate it; reach around sterile surfaces if necessary
• Watch for long hair hanging over sterile surfaces
• Place lids with sterile side DOWN; don't turn lids upside down; nothing will jump up and contaminate the lid
• Use slow, deliberate movements to avoid inadvertant contamination

FINAL NOTE

Accumulated waste materials can pose a contamination hazard. A microbiology laboratory can become inundated with old cultures unless a well organized system for disposal of is in place. Even a few people can produce so much contaminated material, that if teams don't take care of their own materials someone will spend at least a week just cleaning up the place. All cultures must be sterilized before disposal or cleaning of lab ware. To make disposal as efficient as possible, please get rid of materials you no longer need as soon as possible.