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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

A Brief Introduction to Media for Microbiology

This guide introduces methods and general considerations for preparing media for culturing bacteria. You may find the background information to be useful as you plan strategies for isolating and culturing rare and/or "uncultivable" species.

Introduction

Microbiololgy encompasses the study of viruses, bacteria, fungi, and protists, however there is plenty to do just studying bacteria. Bacteria are ubiquitous inabitants of soil, streams, food, on and in us – in virtually all habitable (and some seemingly inhabitable) locations on earth. They can make us wine, yogurt, and garden compost, and without them we couldn't even digest our food. All nitrogen would eventually be lost to the atmosphere without them. Some species have become indispensible tools for research and medicine, supplying us with recombinant DNA, enzymes, and designer drugs, among many other uses.

Bacteria also can make your breath stink, rot your teeth, clog your lungs, give you Montezuma's revenge, and kill you if you (or your physician) are not careful. You have undoubtedly heard of pathogenic Escherichia coli, and "flesh eating" and antibiotic-resistant bacteria. Microbiology has some exciting (perhaps even scary) years ahead of it.

Media

General and specialized media are required for bacterial growth and for characterization. The media you prepare are, in fact, research tools. Peruse this section and use it as a reference as needed. The basic procedures can be applied to almost any type of assay or culture requirement for propagation of obligate aerobes and facultative anaerobes.

Media requirements

Bacteria display a wide range of nutritional and physical requirements for growth including

• Water
• A source of energy
• Sources of carbon, nitrogen, sulfur, phosphorus
• Minerals, e.g., Ca2+, Mg2+, Na+
• Vitamins and growth factors

Microorganisms may be grown in liquid, solid or semisolid media. Liquid media are utilized for growth of large numbers of organisms or for physiological or biochemical studies and assays. Some species, such as Streptococcus or Staphylococcus, often demonstrate typical morphologies only when grown in liquid media. Solid media are useful for observations of characteristic colonies, for isolation of pure cultures and for short-term maintenance of cultures. Usually, the preparation of a solid medium for growth simply includes the addition of 1 to 2% agar to a solution of appropriate nutrients. Agar is a complex carbohydrate extracted from marine algae that solidifies below temperatures of 45šC. It is not a nutritional component.

Frequently, we grow bacteria in complex media because we simply do not know enough about the organism or organisms to define all of their requirements for growth and maintenance. With complex media neither the chemical composition nor the concentration of substrates are defined. These media frequently contain nutrients in the form of extracts or enzymatic digests of meat, milk, plants or yeast. For fastidious organisms we must often use delicious-sounding concoctions such as tomato juice agar or chocolate agar, or something less appetizing (but nutrient-rich) such as brain-heart infusion broth or blood agar.

There is no single medium or set of physical conditions that permits the cultivation of all bacteria, and many species are quite fastidious, requiring specific ranges of pH, osmotic strength, temperature and, in the case of microaerophils, an oxygen content of rather narrow range. Historically, micrbiologists have relied on trial and error to establish suitable growth media. You will culture bacteria on a relatively "lean" medium called Reasoner's 2A agar, a.k.a. R2A agar. R2A agar was developed to reveal the greatest diversity of bacteria in water samples, and thus it is a very good choice for our purposes.

We will use agar plates to isolate and purify cultures and for some assays, as well as for short term maintenance of cultures. We will use agar slant tubes for longer term maintenance of isolates. We use specialized broths (liquid media) to grow isolates for many of our assays. We will prepare most of our agar and broth from pre-mixed dehydrated media.

About dehydrated media

We purchase pre-mixed dehydrated media in the form of granules or powder, and rehydrate the media by mixing a measured amount of medium per measured volume of deionized water. Instructions for rehydration are usually printed on the container (e.g., 18.2 gms/liter for R2A agar). Some media such as phenol red broth or decarboxylase media require that you add a nutrient component and/or adjust pH before sterilization. Some formulations include heat-labile components that must be filter-sterilized rather than steam-sterilized. Watch for special instructions in recipes and on bottles.

R2A agar consists of proteose peptone, caseamino acids, yeast extract, dextrose, starch, and inorganic salts. It is specifically formulated to allow the culturiing of bacteria that would be crowded out by species that grow much faster on richer, more complex media. We may also try culturing isolates on a richer medium called tryptic soy agar (TSA). TSA consists of a pancreatic digest of casein (milk sugar) and a papaic digest of soybean meal, with sodium chloride and agar.

Sterilizing media

When fungal spores or bacteria-laden microscopic particles make contact with your plates, broths, and tubes colonies happily reproduce and your precious media eventually resemble something out of an abandoned full refrigerator. One can't recognize individual colonies when the plates are covered with fuzz! No untreated surface in the lab is sterile, and nearly all dust and other particles have spores or active cells on their surfaces. Obviously, then, all labware and all media must be sterilized before use. We sterilize most media and supplies using a steam autoclave to produce moist heat. Other methods, including filtration, ethylene oxide, radiation, or ultraviolet light, may be necessary if components are heat-labile or materials are not heat-resistant.

An autoclave is designed to deliver steam into a pressure chamber, generating high heat and pressure at the same time. Heating media to above 121 degrees C for a sufficient period of time should destroy all living cells and spores. High pressure (typically 20 lbs/sq. in) allows the temperature to exceed 100 degrees, which can't be accomplished with steam at one atmosphere. 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.

To properly use an autoclave

• Know the instrument - most are now fully programmable, older ones may have manual controls
• Choose the correct program for your materials; we use gravity (fast exhaust with drying) for dry supplies and a liquid cycle (slow exhaust) for agars, broths, and buffers
• Materials may need to be above 121 degrees for 45 min or more to ensure sterilization; the larger the volume the longer it takes for heat to penetrate the interior
• Ensure that the door is closed properly and securely
• Don't relay exclusively on the safety lockout features – ensure that the temperature is well below 100 degrees before attempting to open the door
• Crack the door to allow steam to vent, keeping face and hands well away from the opening
• ***CAUTION*** Exposing tightly stoppered bottles to variable pressures invites explosion and injury. When heating any liquids using any method, take care disturbing the flask or bottle. Material near the bottom may be superheated and boil over when moved. Stoppers, caps, covers, must be vented - never make them fit tightly.

Agar plates

Agar itself has no nutritional value. Agar provides a surface for growth that allows bacteria to take up nutrients that we mix into the agar solution. We use clear plastic disposable petri dishes, typically 95 or 100 mm in diameter, 20 per sleeve. Most dehydrated agar media will contain 1.5% agar (w/v). We prepare and sterilize an agar mix in a flask or bottle, then pour agar into plates either at a lab bench or (preferably) in a sterile cabinet.

Preparing agar plates

When we try to sterilize too large a volume for too short a time we wind up with contaminated agar plates. When we aren't careful mixing agar initially and ensuring that the concentration is uniform following sterilization, we may end up with lumps, or agar that fails to solidify. In our laboratory in the Anderson Biological Laboratory building, Rice University campus, we have exprerienced the fewest issues by following the procedure below.

1. Weigh 18.2 grams of R2A agar (R2A plates) or 40 grams of Trypticase Soy Agar (TSA), place in a clean 2 liter erlenmeyer flask, and add 1 liter of deionized water just before loading the materials into the autoclave. If you don't need that much, you can use 20 gm of TSA and 500 ml of water in a 1 liter erlenmeyer or a 1 liter bottle. As you pour the water in, be sure that it washes down any powder that is clinging to the sides of the flask. Note that containers used for media must have vented tops and should be capable of holding at least 20% more than the intended volume of medium, to allow for expansion during sterilization. A top made of aluminum foil works fine.
2. Swirl the flask vigorously to mix the powder into the water to produce a homogeneous suspension. With TSA it is diffucult to make it completely homogeneous; don't worry about a few small lumps.  Cover the top of the flask with foil, put it in an autoclave tray, and place immediately into the autoclave.
3. Autoclave on an appropriate cycle. We have found that we need a liquid cycle with 45 minute sterilization time for 1L volumes of agar.
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 a 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.

Broth 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. Instead of sterilizing broth media and then distributing it as we do with agar, we prepare broth in a beaker, distribute it into tubes in an autoclaveable rack, cap the tubes, then sterilize it all at once. To mix broth, layer the dehydrated media onto the surface of a measured volume of water in a beaker, use a stir plate and stir bar to mix, and distribute into the tubes using a pipette or syringe. Unless heating is recommended, you should not plan to heat the suspension to dissolve components. Cap your tubes using loose or vented closures, sterilize on a liquid cycle, and allow the media to cool.

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

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.