BISC209/F13: Lab1

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Wellesley College- BISC209 Microbiology- Fall 2013


Lab 1 Introduction to the Microbiology Lab and to the Soil Bacterial Community Project

In this first lab you will learn:

  1. What a microbiologist does
  2. How to sample soil from a habitat without contaminating it to start your semester long project
  3. How to work safely in a microbiology lab by practicing aseptic technique so that you don't contaminate yourself or your cultures
  4. To begin to become familiar with some of the basic equipment and procedures used in microbiological investigation, such as streaking for isolation on various types of media
  5. To use a lab notebook to record the progress of your experiments

Introduction To Microbiology

Welcome to the unseen world of microorganisms. For most of us, microbes are out of sight and out of mind and the human population, by and large, would prefer it that way! Nevertheless, microbes have a major and continuing impact on us and on our planet; therefore, it behooves us to understand them better. Understanding the microbial world is a huge undertaking. A discipline that defines its scope as including all life forms (and some non-life forms like viruses and prions) that are invisible to the unaided human eye has a barely conceivable number of members. As you can imagine, the diversity of such a group is overwhelming.

Where do we begin the study of microbiology? It's good to start with appreciating the power of these tiny, unseen life forms to thrive and spread without our permission or knowledge. It is also wise to recognize that, although only a tiny fraction of the microbes in our world are disease causing, there are devastating infections caused by microbial pathogens. Although none of the microorganisms that we will knowingly work with this semester are common human pathogens, we require that you read and agree to certain rules for working in the microbiology lab that are designed to keep you from infecting yourself, your classmates, and the community. We will also begin to learn aseptic techniques that will reduce the chance of contaminating your cultures or the chance that your cultures will contaminate you.

Lab Safety

Please download, read, and sign your agreement to follow the Wellesley College Microbiology Lab Safety regulations: Media:Safety_rules_micro_3.doc. Bring your signed Safety sheet to the first lab and give it to your instructor. The safety rules can be found in this lab wiki at: BISC209/F13:ReadMe

Please watch these YouTube videos on lab safety at | and |

Introduction To the Tools and Techniques of Microbiology

Whether you are trying to keep a desired organism from being overgrown by a contaminant, or you are attempting to prevent contaminating your soil sample, yourself, your lab bench, or your lab partner with unwanted microorganisms; awareness of potential sources of contamination is critical. Your success in the lab depends on being open to learning and adopting the standard procedures used in microbiology. Today you will practice asepsis when you collect your soil sample and you will practice aseptic transfer technique when you begin to culture your soil sample bacteria.

Asepsis and Aseptic Transfer

Microbiologist must constantly be aware of the ubiquity of microbes on every surface and in the air. We must follow set procedures designed to avoid the inadvertent contamination of microbes from the environment into our samples and avoid contaminating ourselves with our cultures. These procedures are called aseptic technique. You will need to learn them well and follow them rigorously at all times throughout this semester, beginning today.

Your skin is covered with a thick coating of bacteria and eukaryotic microorganisms. It is estimated that every cubic meter of room air contains at least 106 fungal spores and 104bacteria; therefore, touching anything sterile with your hands or any part of your body immediately negates its sterility. If you maintain sterility successfully but you leave sterile samples or equipment uncovered for any longer than absolutely necessary, you have also increased the likelihood that that equipment or sample contains microbes from the air rather than exclusively from the source you desire. When we gather our equipment today and take a soil sample for culture, be aware of the potential sources of contamination and minimize the risk by avoiding touching the part of your equipment that will come in contact with your sample. Work quickly so that your sample is exposed to air or other potential contaminants for as short a time as possible. It is impossible to avoid all sources of contamination but following aseptic technqiues will minimize the risk.

Aseptic Transfer
Please watch the YouTube video on how to use the Bunsen burner |

and the YouTube video on broth to broth Aspetic Transfer Technique |

Manipulation of the many tubes, plates and transfer tools that you will use in each lab requires patience and practice. Mastery is vital to success in the microbiology laboratory. By the end of the semester you will become proficient at many of the transfer methods, you will know when and what to sterilize, and you will be able to keep pre-sterilized tools sterile.

Begin Soil Microbial Communities & Diversity Project

Link to an overview of the Project: BISC209/F13:Project

Soil Sampling

Gather your equipment in teams of 4 before heading out to select a site to sample the soil. Wash your hands. Take off your lab coat and leave it in lab.
Take with you the following equipment for each sampling site (teams of 4):
Your lab notebook and a pen;
1 JMC 18in = 45.7cm steel soil corer with a core diameter of 0.75 inches = 1.9cm (looks like a metal hollow rod with the T at the top)
1 small garden marker to designate where you take your soil sample. Label it: BISC209 Micro Course; date; TUES (or WED) LAB - Soil Sample Habitat Code ______.(Ask your instructor to give you a code to distinguish your habitat)
Also take your kit for each sampling site in a gallon size ZipLock bag containing:
(1) clean spatula or spoon,
(1) 20ml orange top sterile conical bottom plastic tubes labeled with your group's color code, initials, your lab section, the date, the greenhouse room that you plan to sample, and the code letter your instructor has given you to distinguish your soil sample from your classmates'.
(1) black Sharpie,
(several) paper towels
(4) pairs of disposable gloves in the appropriate size for you and your teammates

With your teammates, go to the selected habitat you are to investigate. Before you take your soil sample, look around at this habitat and choose a location to sample. Record your impressions as you discuss the habitat with your group. Does it seem cool, warm, or cold; dry, moist, or average in humidity? Is there abundant sun, shade, or is it mixed? What else strikes you about this environment that you may want to add to your notes? Today you will draw a scaled "map" in your lab notebook to show where you sampled, containing enough detail so that someone else could easily locate the area even if the marker you will place here is removed.

Avoid very wet soil and highly compacted soil but get close to a plant and record its name.

When you and your teammates have agreed on where to take your soil sample, get your equipment laid out and ready. Place 2 or 3 paper towels down end to end in an open area near where you are going to sample and put on your gloves. The gloves are not sterile but be careful to avoid contaminating the exterior surface of the gloves with skin flora by touching your skin or anything other than the soil area you are going to sample.

To sample the soil, brush away any leaf debris or non-soil material that might end up in your sample.
Push the soil sampling device straight down, putting force with both of your hands until your corer is in about 3-4 inches. It is best to twist the corer straight into the ground.

Empty the corer on the paper towels you made ready nearby. Knock the side of the corer and the soil should emerge as an intact cylinder of soil. If it doesn't, you may pull it out with your gloved hands. It doesn't matter if you don't get an intact cylinder. Your goal is to sample equally from this soil from the lower, middle, and upper areas of the core (avoiding the very top 30mm [1 inch] of surface soil), collecting enough soil to fill the 20ml tube abut 2/3 to 3/4 full.

Spoon (you may use the spatula you brought) the soil sample into the labeled 20ml conical tube. Try to make the area look undisturbed. Mark the spot sampled by the team with your labeled garden marker.

Return as quickly as you can to the lab with your labeled soil sample tube and all of your equipment. We have a lot to do today with these samples to start our analysis of the bacteria in this soil community.


Soil Preparation

Sieve the fresh soil sample using the sterile 2 mm pore size sieve, beaker and pestle. Wear gloves to avoid contaminating the soil with human microbes. Save the sieved soil in the covered beaker.

Soil Extract Preparation

Weigh 1 gram of sieved soil using the top loading balance and add it to 100 mL of sterile water with 0.0001% Tween 80 (a surfactant that helps remove bacterial cells from the soil particles they tend to cling to). You will find the water/Tween mixture premeasured for you in a sterile 250 ml flask on your bench. Swirl to mix--- don't add a magnetic stir bar yet. Pour this soil suspension into a clean blender jar. Be careful not to contaminate the inside of the lid with your fingers when you place the lid on the jar. Blend at highest speed for 3 pulses of 10 seconds on and 10 seconds off. Pour all of the suspension back into the 250 ml flask and add a sterile magnetic stir bar (on your bench). Place the flask on a magnetic stirrer at medium speed and mix for at least 15 min. Stop the stirring and let the soil settle until the larger particulate matter settles to the bottom. (Not all visible particles need to settle, just the big stuff.) When you use it, be sure to take your aliquot from the top portion that lacks the big particulate matter. In making this extract, you have created a 10-2 or 1/100 dilution, which can be described in concentration units as 1% (wt/vol) since there is 1 gram/100ml. Save the extract!

Drying Soil for Calculation of DRY weight

Weigh three 1 gram samples of your mixed and sieved soil into properly labeled aluminum weighing boats (label each with a piece of your team tape color on which you have identified the soil sample). You will find the aluminum weigh boats near the scale. It is important to use the aluminum boats and not the plastic ones so the soil can be oven dried. Record the weight of the aluminum weighing boats in your lab notebook before you tare or zero the balance and add the 1 gram of soil. Leave the weighed samples on the metal tray marked with your lab section found beside the top-loading balances. These soil samples will be oven dried at at a low temperature (70°C) for you and returned to you for use in Lab 2. Drying the soil necessary for our enumeration experiments because we want to calculate the number of microbes per gram of DRY soil. This is a more standardized calculation since soils vary in the moisture content.

Assessing the Number of Microbes In Your Soil (in pairs)

After preparing the 1% soil extract, please label a sterile 1.5 mL microcentrifuge tubes with your work-site letter using a piece of your team color tape and an indelible Sharpie. Each site needs to prepare 2 tubes. Make sure all the labeling is legible! Aseptically transfer 1.0 ml of your soil extract to these tubes using your P1000 micropipet and different sterile tips. If you are inexperienced at using a micropipet, ask your instructor for assistance. Be careful not to contaminate the pipet tips by touching them to anything including your hands.

4'-6-Diamidino-2-phenylindole (DAPI) aqueous solution Stock conc. is 1mg/ml: DAPI is known to form fluorescent complexes with natural double-stranded DNA.

If the microfuge samples will not be processed for a while: You need to add paraformaldahyde at an effective concentration of 4% to preserve the microbial cells in your soil extract samples. Ask your instructor about this step.

  1. Add 5μL of 1mg/ml DAPI stock (Thermo scientific prod. # 62248) stain to 1mL of a fresh 1% diluted soil extract so that the effective concentration of DAPI is 5 μg/mL.
  2. Incubate at 4°C for 20 min in the dark.
  3. Set up a vacuum flask and filter apparatus (125 ml side-arm flask, Borosilicate base and fritted glass filter support with rubber stopper) in the basket so that the flask is level.
  4. Connect the rubber hose to the vacuum jet.
  5. Carefully place a 2.5 cm diameter (Whatman GF/A) 1820-025 (0.45 µm pore size) backing filter smooth side up and a 25 mm diameter Whatman 110656 black polycarbonate (nucleopore) 0.2 pore size filter onto the fritted glass filter support shiny side up.
  6. Add the borosilicate glass funnel onto the base and clamp the two sections together using an anodized aluminum spring clamp.
  7. Apply 1 mL sterile deionized water to the surface of the filter, turn on the vacuum full force. Wait until all the water is removed.
  8. Turn off the vacuum jet.
  9. Remove the rubber hose from the vacuum jet nozzle to dissipate the remaining vacuum.
  10. Replace the hose on the vacuum jet.
  11. Repeat steps 7-10 so that you have rinsed the filter and equipment twice.
  12. Be sure the flask sits level in the basket: Hold it level if necessary.
  13. Transfer all of the DAPI stained extract (made in step 1) onto the filters.
  14. Turn on the vacuum full force.
  15. Once the sample is completely filtered, rinse with 1 ml aliquots of sterile water twice to remove any excess DAPI.
  16. Turn off the vacuum jet.
  17. Remove the rubber hose from the vacuum jet nozzle to dissipate the remaining vacuum.
  18. Use a marker to label the left side of a glass slide with your site ID, your initials, and T (for Tues lab) or W (for Wed. lab).
  19. Take the apparatus apart by removing the clamp. Be careful with the glass funnel.
  20. Use forceps to carefully remove the top Black Polycarbonate Filter and place that filter on the labeled microscope slide. Discard the PTFE filter.
  21. Apply 1 drop of Fluoro-Gel with Tris buffer (Electron Microscopy Sciences Cat #17985) to the top of the filter.
  22. Place a coverslip over the filter.
  23. Place your slide flat in the slide box and close the box cover.
  24. Your instructor will place the slide box in the fridge until she can use the Nikon Eclipse 80i fluorescent microcope in L318C to view and photograph the labeled DNA at 1000x magnification. Since we only have one microscope, your instructor will prepare the photographs for you. She will use the Nikon NIS-Elements Imaging software, to photograph a representative field of view in white light to see relative size and shape of the organisms in the sample. (Keep in mind that not all of the DNA will be from bacteria: some will be eukaryotes'.). After saving this image she will then photograph the same field using the UV light at 350nm excitation wavelength (filter #1) so you can visualize the fluorescent labeled DNA.
  25. All the images will be saved to a designated file folder and provided to you.
  26. In our next lab you will count the unique spots of blue fluorescence; each indicating a soil microbe's genetic material (bacterial chromosome or eukaryotic nucleus). It is assumed that the microbes were arrayed in a Poisson distribution when the vacuum distributed them on the filter when the liquid was removed. If one wanted to publish this work, 20 fields or ~400 bacteria would be enumerated in order to have an acceptable estimate of the number of bacteria per ml.
  27. We will pool the data for each experimental site and calculate our estimate of the average number of microbes per gram of soil next week.


To perform a plate count of your culturable soil sample microorganisms you must first dilute the soil serially so that you will reach a dilution where have a countable number of colony forming cells spread out over the plate. The goal is to obtain 30-300 well isolated colonies at one of these dilutions. If you don't remember how to make a serial dilution, here is a link to a helpful animation for making dilutions

Standard Plate Count of Soil Microorganisms on Dilute Nutrient Agar
Do this test in pairs for your soil sampling group so that there are 4 replicates for each site.
The Soil Extract you prepared is a 1:100 soil dilution (1 gram/100 ml). This could also be called a 1% (w/v)suspension.
Gather the following materials to start your quantitation:
-5 sterile 13 x 100 mm size sterile glass tubes with caps
-Sterile disposable plastic individually wrapped 1 ml pipets and pipet pump
-10 sterile dilute (1:10 diluted from full strength) nutrient agar plates,
-10 sterile plastic disposable spreaders

Setting Up a Standard Plate Count: (pairs)

  1. Label a set of five 13 mm glass tubes 10-3, 10-4, etc. ---through 10-7.
  2. Label the destination plates of dilute nutrient agar: two plates for each dilution. Include your name, date, lab section, soil sample code letter, and the dilution.
  3. Slightly dehydrate the medium on all of the plates. To do this you will turn on the fan on one of the laminar flow hoods; clean the surface with alcohol; place all the plates in the hood; position the covers so they are slightly ajar; leave them for 10 minutes or until the medium surface shows no visible moisture. Replace the tops and bring them back to your bench.
  4. Use a sterile 1ml disposable pipet and your blue Pipetman to pipet 0.9 ml of sterile water into the 5 tubes labeled in step 1. (You may use the same sterile 1ml pipet for all tubes.)
  5. (If you aren't very confident that you know how to use a micropipet properly or you want a review, ask your instructor for a quick tutorial before you start this step.) Using your P200 micropipet and autoclaved tips, transfer 100 microliters (0.1ml) of your soil extract (1:100 dilution) to the tube labeled 10-3; mix well by vortexing.
  6. Using a new tip, transfer 100μL (0.1ml) of the 10-3 dilution to the tube labeled 10-4. Mix well by vortexing. Mixing 0.1 ml of the 10-3 dilution with 0.9ml of sterile water makes a 10-4 dilution.
  7. Continue to transfer 100μL (0.1ml) aliquots (after mixing well) from each dilution to the next tube of 0.9 mL water until you have carried the dilution to 10-7. Use a new tip for each transfer.
  8. Starting with the most dilute extract and a new tip, transfer 100μL (0.1ml) and dispense it to the center of the first pre-labeled culture plates of dNA for that dilution. Use a sterile plastic disposable spreader to gently push the dispensed sample two or three times clockwise around the dish, and then several times counterclockwise. Make sure all of the surface area of the plate has been inoculated. Don't press too hard as force will cause the microorganisms to collect at the edge of the spreader, resulting in uneven distribution.
  9. Repeat step 8 to inoculate the second plate for that dilution.
  10. Repeat steps 7, 8 and 9 for the rest of your dilute nutrient agar plates. You should end with each of the five dilutions on two dilute nutrient agar plates for a total of 10 plates/pair.
  11. Allow the moisture to be absorbed into the agar before inverting the plates. Put a labeled piece of your team color tape around each set of plates (separate the two media). Microorganisms are usually cultured on solid medium up-side down to avoid condensation issues from temperature changes. Incubate your cultures at room temperature (RT) in a rack designated by your instructor until next week.


1. Culture plates, stocks, etc. that you are not finished with should be identifiable with your team color tape and well labeled. Place the labeled cultures in your lab section's designated area in the incubator, the walk-in cold room, or at room temp. in a labeled rack. If you have a stack of plates, wrap a piece of labeled team color tape around the whole stack.

2. All culture plates that you are finished with should be discarded in the big orange autoclave bag near the sink next to the instructor table. Ask your instructor whether or not to save provided stock cultures.

3. Remove tape from all liquid cultures in glass tubes and place the glass tubes in racks by the sink near the instructor's table. Do not discard the contents of the tubes. Place the non-disposable caps for these tubes in the wire basket provided in the clean-up area near the sink.

4. Glass slides or non contaminated and empty disposable glass tubes can be discarded in the glass disposal box.

5. Make sure all contaminated, plastic, disposable, serologic pipets and used contaminated micropipet tips are in the small orange autoclave bag sitting in the plastic container on your bench.

6. If you used the microscope, clean the lenses of the microscope with lens paper, being very careful NOT to get oil residue on any of the objectives other than the oil immersion 110x objective. Move the lowest power objective into the locked viewing position, turn off the light source, wind the power cord, and cover the microscope with its dust cover before replacing the microscope in the cabinet.

7. If you used it, rinse your staining tray and leave it upside down on paper towels next to your sink.

8. Turn off the gas and remove the tube from the nozzle. Place your bunsen burner and tube in your large drawer.

9. Place all your equipment (loop, striker, sharpie, etc) including your microfuge rack, your micropipets and your micropipet tips in your small or large drawer.

10. Discard any remaining sieved soil. Return it to the original 50 ml conical tube and dispose of it in the trashcan (NOT the autoclave bag).

11. Discard the remaining soil extract.

12. Move your notebook and lab manual so that you can disinfect your bench thoroughly.

13. Take off your lab coat and store it in the blue cabinet with your microscope.

14. Wash your hands.

15. See you next time!


Graded Assignment: NO graded assignment this week:)

Do before next lab:
Make sure you have read all of the introductory information on the wiki about the project on the project home page BISC209/F13:Project, reviewed what was accomplished in Lab 1, and read Lab 2 carefully before coming to lab next time. Prepare for Lab 2 by making brief flow diagrams of the work you will do in the appropriate sections of your notebook. There are some helpful hints about organizing your lab notebook in the BISC209/F13:Resources section of the wiki.