BIOL 302 Lab schedule, Fall 2022:
LAB
DATE
1
Tue 8-23
2
Thu 8-25
3
Tue 8-30
4
Thu 9-1
5
Tue 9-6
6
Thu 9-8
7
Tue 9-13
8
Thu 9-15
9
Tue 9-20
10
Thu 9-22
11
Tue 9-27
12
Thu 9-29
ACTIVITIES
Lab safety
Introduction to the course
Aseptic technique
Isolation of microbes from the environment
Microscopes and Kohler Illumination
Microscopes and Kohler Illumination, cont’d
Using Micropipettes
Pure culture techniques
Principles of staining and Simple stain
Winogradsky column
Capsule stain
Gram stain
Dilutions
Acid-fast stain
Live/Dead stain & fluorescence microscopy
Direct counts
Media
Anaerobic culture techniques
Maintaining cultures
Sterilization procedures overview
Quantification of bacteria
Endospore stain
Bacterial identification virtual lab
Growth Curve
Bacterial Motility
Plan group project and perform reference
search (primary vs. secondary literature)
Quorum sensing
Dichotomous key: Creating a key for
identification
Biochemical reactions (Sugar Fermentation
tubes and MR-VP)
QUIZ OR ASSIGNMENT
Bring Winogradsky column
materials;
First notebook check will occur
anytime 8-30 through 9-15
Quiz (Sessions 1-4);
Dilutions practice 1
Dilutions practice 2
Dilutions practice 3;
Second notebook check will
occur anytime 9-20 through
10-13
Quiz (Sessions 5-9); Skills test
deadline
Dilutions test; First group
project plan and materials
request form due end of class
Bring environmental sample;
Dichotomous Key due
Continue Biochemical activities (Amylase,
Catalase, Oxidase)
Group projects: Environmental sample
extraction
13 Tue 10-4
Continue Biochemical activities (Indole,
Citrate, TSI, SIM)
Group projects
Wed 10-5: Lab written exam 1 (during lecture); covers material through Lab Session 11
14 Thu 10-6
Lab Practicum (during lab)
Continue group projects
15 Tue 10-11
Plagiarism exercise
Continue group projects
16
Thu 10-13
17
Tue 10-18
18
Thu 10-20
19
Tue 10-25
20
Thu 10-27
21
Tue 11-1
22
Thu 11-3
23
Tue 11-8
24
Thu 11-10
Complete group projects
Prepare poster
Group Project Poster Presentations
Biofilm lab
Ames test
qRT-PCR assignment
Plan second group project and perform
reference search
Biofilm results
Ames test results
Plan second group project and perform
reference search
Plaque Counts
Kirby Bauer
Plaque Count results
Kirby Bauer results
Epidemiology lab
Winogradsky column results
Third notebook check will
occur anytime 10-18 through
11-15
Bring Ames Test sample
Quiz (Sessions 12-18);
Second group project plan due
at end of class
qRT-PCR assignment due
Second group project plan
deadline (end of class)
Epidemiology lab results
Group project
Termite gut symbionts
Quiz (Sessions 19-23)
Eukaryotic microbes
Continue group project
25 Tue 11-15
Continue group project
Wed 11-16: Lab written exam 2 (during lecture); cumulative but focuses on material from
Lab sessions 12-24
26 Thu 11-17
Lab practicum (during lab)
Continue group project
Tue 11-22 and Thu 11-24 Fall recess; no lab
27 Tue 11-29
Continue group project
28 Thu 12-1
Continue group project
29 Tue 12-6
Continue group project
Draft of paper due; bring 4
Peer review of group paper
copies for peer review
30 Thu 12-8
Powerpoint Presentations
12-12 by 5pm: Final paper
(see below)
Final Group Paper: A hard copy must be turned in to your lab instructor’s mailbox (Bio
department office, MH-205) AND an electronic copy submitted to the Turnitin link in Canvas NO
LATER THAN 5:00pm on Monday December 12. NO E-mailed papers.
BIOLOGY 302L GENERAL MICROBIOLOGY LAB
(Edited Fall 2022)
Welcome!! This lab course complements the General Microbiology Lecture by providing handson experience working with microbes. The course begins with basic microbiological lab
techniques, such as aseptic technique, staining, bright field microscopy, pure culture techniques,
and dilutions. The rest of the semester applies the basic techniques to investigate the
characteristics of microbes. There are two group projects designing “open discovery” studies to
identify and characterize microbes in environmental samples.
Each student will be assigned five slides and a lab drawer for the semester. Clean your drawer
with disinfectant before you put in any materials. Each student must supply their own Sharpie
and Lab coat. Personal belongings are not to be stored in the laboratory.
To set yourself up for success in BIOL 302 Lab:
Before class
• Check which activities are scheduled for that day (see schedule).
o For example, for Session 2: Microscopes and Kohler Illumination cont’d, Using
Micropipettes, Pure culture techniques, Principles of staining and Simple stain
• Read the lab manual pages for those activities.
o For additional preparation on performing lab techniques, view the lab video for each
activity (links on Canvas).
• Complete your pre-lab: write the Date, Title, Purpose, and Materials & Methods for each lab
activity in your lab notebook.
o The pre-lab will be checked when you enter class each day. It is best to paraphrase
and use your own words when you can, rather than copying everything directly from
the lab manual, since the purpose of the pre-lab is to prepare you to complete the lab
successfully. If you are starting several different activities during a single class
period, place each on a separate page and leave enough space to record all
information for each experiment.
During class
• Practice understanding of the material during class discussions and Q&A with your
instructor.
• Carry out lab activities
• Complete notebook entries (Results, Discussion, and note any modifications to the Methods)
for that day’s activities before the end of the lab period, while your TA is available for
questions. See the “Lab Notebook” section of the lab manual for more details.
1
TABLE OF CONTENTS
Lab safety........................................................................................................................................3
Laboratory notebook .....................................................................................................................4
Aseptic technique ...........................................................................................................................7
Isolation of microbes from the environment ...............................................................................9
Microscopes ..................................................................................................................................11
Kohler illumination......................................................................................................................18
Using micropipettes .....................................................................................................................23
Pure culture techniques ...............................................................................................................25
Principles of staining and Simple staining .................................................................................29
Winogradsky column ...................................................................................................................33
Capsule stain ................................................................................................................................35
Gram stain ....................................................................................................................................37
Working with dilutions ................................................................................................................40
Acid-fast stain ...............................................................................................................................42
Live/dead stain .............................................................................................................................44
Direct counts .................................................................................................................................46
Media.............................................................................................................................................48
Anaerobic culture techniques .....................................................................................................52
Maintaining cultures....................................................................................................................55
Sterilization...................................................................................................................................56
Quantification of bacteria ...........................................................................................................57
Endospore stain ............................................................................................................................62
Bacterial Identification Virtual Lab...........................................................................................65
Growth Curve...............................................................................................................................67
Bacterial motility ..........................................................................................................................70
First group project: Investigation of an environmental sample ..............................................74
Quorum sensing ...........................................................................................................................78
Dichotomous key ..........................................................................................................................81
Biochemical reactions ..................................................................................................................82
Summary of biochemical reactions ............................................................................................93
Biofilm formation .........................................................................................................................97
Ames test for mutagenicity ........................................................................................................100
Plaque counts..............................................................................................................................103
Kirby-Bauer: antibiotic susceptibility testing .........................................................................105
Epidemiology ..............................................................................................................................109
Termite gut symbionts ...............................................................................................................115
Eukaryotic microbes ..................................................................................................................118
Second group project .................................................................................................................121
2
LAB SAFETY
Safety precautions must be followed whenever working in the microbiology lab.
1) Many microorganisms are potential pathogens. Cultures must be handled with care and
disposed of properly.
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Always wash your hands after working with microbes and before leaving the lab.
Do not sniff cultures or pipette by mouth.
If there is a spill, wipe up the material using a disinfectant-soaked towel.
Place any materials with live organisms, including paper towels and other disposable
items, into the appropriately marked containers.
Label all plates and tubes clearly. Keep them closed and stored properly when not in
use.
Avoid creating aerosols when flaming your loop.
Wipe your work surface with disinfectant when you are done with your work.
2) Wear proper personal protective equipment. Lab coats and closed-toe shoes are required
in the lab at all times. Gloves are required when conducting experiments. Eye protection
(glasses or goggles) is required for any experiments involving liquids.
3) Place the bunsen burner in a safe position when in use and turn it off when not in use. To
prevent accidents, long hair should be tied back and all unnecessary books should be
stored.
4) Keep dyes and other potentially harmful chemicals securely closed and in their assigned
storage sites when not in use. Some dyes stain the skin and clothing and certain
chemicals can make holes in clothing.
5) Check the temperature of the water baths by reading the thermometer before use.
6) Don't try to hold or carry too many items at once. Take care when working with hot
materials. Do not carry tubes or bottles by their lids.
7) Dispose of broken glass appropriately. All sharps must be deposited in the sharps
containers.
8) Know the location of safety equipment. The lab is equipped with an eye wash station,
first aid kit, and shower.
Above all, if you are uncertain of a procedure or how to handle the equipment, ask your
instructor.
3
LABORATORY NOTEBOOK
Keeping an accurate and precise lab notebook is a critical skill for scientists. Your notebook is a
useful and valuable document; it should be detailed enough to allow you or others to perform
each laboratory activity, and to help you troubleshoot when results are unexpected. Scientists
keep their lab notebooks up to date in real time as they perform experiments; the lab
notebook is NOT a lab report with a specific due date.
When writing in your lab notebook, more is not better! Strive to be precise and complete
but also CONCISE. Think carefully about what you write, and write thoughtfully without
rambling and copying.
Your lab notebook should be kept in a carbon-copy notebook, which can be purchased at Titan
Shop or online. A previous class’s carbon-copy notebook that still has blank pages is acceptable.
The notebook should be organized by experiment, not strictly by date. For an experiment that is
started on one day and completed the next, the notebook entries should be included under the
same experiment. If pages are not pre-numbered, you should number the pages consecutively.
Leave the first few pages blank to serve as a Table of Contents (TOC). List all experiments in the
TOC in chronological order using the following columns: Date, Experiment Title, Page #.
You must bring your notebook to every lab and record observations directly into the notebook
as experiments are performed. It is more important that your notebook is a complete record of
what you did than being spotlessly neat like a final paper. Do NOT write results on scraps of
paper to be re-copied at a later time.
Each notebook entry should include the following sections:
Completed prior to class (the pre-lab will be checked upon entering)
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Title: The title must be descriptive and can come from the lab manual. The title needs to
be written on the top of the page and in the TOC.
Purpose: A brief 1-2 sentence explanation of the aim or purpose of the lab activity.
Materials: A list of all materials used to complete the experiment including positive and
negative controls, equipment, and organisms. (from the lab manual)
Methods: A concise step-by-step procedure for the experiment including specific times,
temperatures, volumes, etc. The Methods in your notebook should be paraphrased/written
in your own words, but be detailed enough that they allow you to perform the whole
experiment without referring to the lab manual (and allow anyone else to repeat the
experiment). You can display the procedure as a flowchart or list.
• Note: If you make any modifications to the procedure while performing the lab, you
must record those modifications in the notebook during the lab.
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Completed during class when results are available
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Results should be recorded in the lab notebook the day they are available. All results
need to be detailed and clearly organized.
o All results figures, drawings or pictures must be labelled with the following
information, if applicable: species name, total magnification, media, etc.
Sketches, particularly with colored pens or pencils, are completely acceptable.
You don’t have to be an artist, you just want to record your observations clearly.
You may choose to take photos of the results but, if you do so, ensure they are
printed and taped into the lab notebook in a timely manner since the notebook
checks are random. Having the photos on your phone does not count. If photos
are printed in black and white, make sure to use words to describe what colors and
details you observed.
o Using words, write down all observations: color changes, growth or no growth,
contamination, # of CFUs, calculations, concentrations, etc. Where appropriate,
organize data into tables.
o For the skills tests, the notebook entry of your results should be from your FIRST
attempt. Results do not need to be perfect; good or bad they ALL go into the
notebook – this applies for all experiments.
Discussion: What can you conclude from your results?
o BRIEFLY summarize then interpret your results. What do the results mean? What
is their significance? (see examples below)
o Provide possible explanations for any unexpected results. List any sources of error
and explain how they may have influenced your results.
Discussion examples:
After gram staining, S. aureus cells appeared purple and spherical. Therefore, S. aureus
bacteria are gram positive cocci.
After incubation of E. coli in Glucose fermentation broth, the culture was turbid, the media
color changed from red to yellow, and there is gas in the Durham tube. Thus, E. coli ferments
glucose to produce acidic fermentation products and gas.
Note: Post-lab questions do NOT need to be answered in the lab notebook. These questions are
for further thought and may be useful when reviewing for exams.
Group Projects: Group project objectives, methods, and results should also be included in the
lab notebook. If you are working on multiple experiments, make sure to anticipate and leave
enough pages to record all information.
5
LABORATORY TOUR
WHERE THINGS ARE IN THE LAB
During the semester you will be using many different supplies and several pieces of equipment.
You will also be generating much material that you will have to dispose of properly. During the
first laboratory period, your lab instructor will orient you to the classroom and to appropriate
safety procedures when working with microbes.
Make sure that you understand where the following items are located:
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Microscopes
Temperature-controlled equipment
§ Incubators (Each section has an assigned space in a 37°C incubator)
§ Refrigerators
• for storing cultures
• for media storage
§ Water baths
Stain/chemical storage
Water for dilutions
Sterile test tubes
Test tube racks
Sterile pipettes
Reference books
Disposal areas (biohazards)
§ contaminated reusable glassware
§ broken glassware
§ cultures
§ plastic plates and other nonreusable materials
§ sharps
Spectrophotometers
Typically, media requested for each experiment will be placed on the rear countertop or in the
water baths. Control cultures will be supplied to each group and the group will be responsible
for their maintenance. Special cultures will be placed in racks on the lab benches when needed.
These cultures will be shared so be careful not to contaminate them.
The laboratory should be found clean and should be left clean. Microscopes must be maintained
properly. If you should find anything out of order or dirty, please report this to your instructor
promptly.
6
ASEPTIC TECHNIQUE
Most of the exercises and experiments you will perform during this semester use sterile media
and pure cultures (progeny of a single organism). We will isolate pure cultures during the next
lab period, but practice aseptic technique today. A series of operations have been developed to
limit the risk of contaminating these materials during manipulations. These operations are
known as aseptic techniques. They also help protect investigators from infecting themselves or
releasing the organisms into the environment. Learn the following rules and procedures and
understand how each requirement contributes to maintaining asepsis. Aseptic technique is an
important skill that should be mastered by all scientists that work with cell cultures.
OBJECTIVES: Students should be able to
1. Describe and explain the principles of aseptic technique.
2. Work with media and cultures without contamination or cross-contamination.
General Rules
1. Work on a clean tabletop. Put all unnecessary items away.
2. Wear a lab coat; wash hands before performing any manipulations and after you are
through.
3. Disinfect the benchtop with an appropriate disinfectant before you begin working and
after you are through.
4. Focus on your work and avoid being disturbed.
5. Keep all bottles, tubes, petri dishes, etc closed as much as possible. Keep tubes and
bottles upright to avoid spills and contamination.
6. Flame bottle and tube mouths before and after use.
7. Sterilize inoculating loops and needles before and after use.
Aseptic Technique for Preparation of Agar Plates
1. Molten sterilized agar medium is available in a water bath at 45-50°C. Make sure that
there is enough water to cover the agar medium. If necessary, use a "doughnut" to
prevent the agar-containing vessel from tipping.
2. Place sterile petri plates on a disinfected benchtop.
3. Remove the lid from the agar-containing vessel and pass the mouth through a flame to
destroy any contaminating organisms. Hold the container at an angle and not vertical.
4. Pour the agar into the petri dishes. Gently swirl the plate to distribute the agar to cover
the bottom of the plate.
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5. If any agar remains in the container, pass the mouth through a flame and close the vessel.
Return it immediately to the water bath.
6. Allow the plates to solidify. Label the plates with the type of media they contain and the
date they were poured.
7. If plates are being stored for more than 48 hours, seal them in a plastic bag.
Aseptic Technique for Transfer of Organisms
a. Microbes can be removed from broth with an inoculating loop or a pipette; they can be
removed from solid media with an inoculating loop or needle.
b. Inoculating loops and needles are made from metal wire and can be sterilized by flaming
until red-hot. The entire length of the metal wire must be sterilized. This is done by
flaming/heating the wire from the handle end to the tip to incinerate all organisms that are
present. Once cooled, the loop or needle can be used to transfer a bacterial sample. After
transferring a sample, flame the loop or needle immediately, before transferring another
sample or putting the loop or needle down on the bench.
c. Flame the mouth of tubes or flasks after removing the lid. Hold the tube or flask at an
angle while transferring a sample. Flame the mouth again before replacing the cover.
d. Dispose of all contaminated pipettes or tips in the appropriate containers.
Practice Aseptic Technique
MATERIALS
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Sterile petri dishes
Melted TSA medium at 50°C
Inoculating Loops
Sterile water
TSB
Pour two TSA plates using the procedure described above. Also, inoculate a loopful of sterile
water into a TSB tube.
Since neither the inoculating implements nor the vessels are to be put down during the
manipulations, the most difficult part of these procedures often is in one person handling
everything simultaneously. Your instructor will demonstrate how this can be done, and you
will practice the technique.
8
ISOLATION OF MICROBES FROM THE ENVIRONMENT
Objectives: Students should be able to:
Isolate microbes from the environment
Correctly label a petri dish
Observe microbial growth on solid media
1.
2.
3.
Only a few environments are sterile, free from living organisms. The most widespread forms of
life are the microbes, many of which can live where larger organisms cannot survive. Some
microbes are composed of eukaryotic cells; they have a true nucleus and membrane-bound
organelles. Other microbes are prokaryotic, a cell characterized by its DNA condensed in a
region called the nucleoid. Prokaryotes are renowned for their metabolic diversity and an ability
to secure a habitat in highly unfavorable conditions. The distribution of microbes based on cell
structure is shown below.
Eukaryotes (True nucleus)
Algae
Fungi
Prokaryotes (Nucleoid)
Protozoa
Bacteria
Archaea
Molecular analyses have revealed that all organisms can be assigned to one of three Domains
(the highest level of classification) based on DNA comparisons. All organisms comprised of
eukaryotic cells belong in the Domain Eukarya. Archaea and Bacteria are two distinct
Domains containing only microorganisms. Eukarya and Archaea are more closely related to
each other than to Bacteria. The "typical" prokaryote that you will encounter in your
environment is a member of the Domain Bacteria.
During this laboratory period, the class will attempt to collect microbes from a variety of
sources. You will be using a solid medium called trypticase soy agar (TSA) that supports the
growth of many organisms.
MATERIALS
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1 Trypticase soy agar (TSA) plate
Sterile cotton-tipped swabs; sterile toothpicks
Sterile water
9
PROCEDURE
1. Expose the agar surface to the environment as suggested below.
2. Using a water resistant marker, the bottom (side with the agar) of the petri plate with:
• your name
• the date
• the environmental exposure.
• the type of media
3. Incubate the plate inverted at 37°C for 48 hours.
Suggested Exposures
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Remove the lid and expose to air in laboratory, hallway, outdoors, or other rooms. Record
the time that the agar was exposed.
Place any material on agar surface (dust, money, candy).
Touch the agar surface, cough on it or comb your hair over it.
Moisten a swab and apply it to a surface. Streak the moist swab over the agar.
Clean between your teeth with a toothpick and spread on the agar.
Use your imagination (hopefully it's not sterile).
RESULTS
Examine your plates and describe or diagram (in your notebook) the extent and appearance of
growth. Include the following information:
• Exposure site
• Total Number of Colonies
• Number of Different Colonies
• Description of Colonies
Post-lab Questions
1. Do you expect all the different microbes, from the environment you sampled, to grow on your
plate, or will some be missing? Justify your answer.
2. We incubated the plates at 37°C; was that the best temperature to use for your inoculum?
Why or why not?
10
MICROSCOPES AND KOHLER ILLUMINATION
OBJECTIVES: Students should be able to
1. Explain the principles of microscopy
a. Differentiate between resolution and magnification
b. Determine the resolving power of a lens
c. Explain the increased resolution of the oil immersion lens
d. Define parfocal, working distance
2. Identify the parts of a brightfield microscope and explain their function
3. Convert measurements between millimeters and micrometers
4. Correctly use and clean the laboratory microscope
The scientific method requires evidence for the determination of facts. If our senses are
limited, we need tools to increase the power and accuracy of our observations and, thus,
our knowledge of the world. For biologists, light and electron microscopes have become
indispensible as "eyes" for discovery about the detailed structure of cells. The purpose of
Kohler illumination, even illumination and high contrast without light artifacts, is to maximize
resolution, the detail provided by magnification.
Resolution is the ability of a viewing instrument (the eye, a camera, a microscope)
to distinguish adjacent objects as separate rather than as one larger object. For example,
how close can two lines get before you will no longer be able to see both of them? How
thin can a single line become and still be visible?
When at close focus the lens of the human eye has a resolving power of approximately 100 µm.
This means that the eye will see two objects as distinct if the space between them is at least 100
µm. If the space is 90 µm, they will appear as a single object. The best light microscopes have
resolving powers of about 200 nm. They will "see" objects or spaces between objects as small as
200 nm. If you look at the period at the end of this sentence you will see a dot. Imagine that it is
a cell composed of many parts. If you magnified the dot 100 fold without increasing resolution
(as in photographs) you would have a larger image but not greater detail. (This is called empty
magnification.) Magnifying the dot 100 fold with a microscope, however, resolves ultra-fine
structures of the image. Thus, the useful magnification of a viewing system depends on its
resolving power.
The two most important factors that determine resolving power are the quality of the lens
system and the wavelength of the source of illumination. Resolution varies inversely with
wavelength; a shorter wavelength allows for greater resolution.
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Light Microscopes
Standard laboratory microscopes are compound microscopes, constructed of two lenses in series
that focus rays of light to illuminate the object. The total magnification is obtained by
multiplying the magnification of both lenses. High quality lenses give light microscopes
resolving powers of about 200 nanometers, a 500- fold increase over that of the eye. This allows
the observer to visualize the outlines of bacterial cells and only glimpse at the larger internal
structures in cells of plants and animals.
Brightfield Microscope
The most common light microscope is a brightfield microscope - one that uses visible
light to illuminate the specimen. You will be required to identify all of these parts (diagram,
next page) and know their functions.
Oil Immersion
As previously stated, resolving power or resolution (r) varies with wavelength of light and
quality (light gathering capacity or numerical aperture (NA)) of a lens.
r = Wavelength/ 2 x NA
The brightfield microscope often uses visible light with a wavelength of 550 nm. The NA of a
low power objective is around 0.25. Thus, the best possible resolution (minimum distance
between resolvable objects) through that lens is 1100 nm.
To increase resolution, either the wavelength of light transmitted through the specimen must be
decreased or the NA of the objective must be increased. Many brightfield microscopes use a
blue filter over the light source to reduce the wavelength of transmitted light to 475nm.
NA depends on the configuration of the lens and the material between the lens and the light
source. Some light is lost when it passes from the specimen on a glass slide through air to the
objective lens. This occurs because the refractive index (ability to bend light) of glass and air
differ. The air space can be replaced by a drop of immersion oil, which has a higher refractive
index, closer to that of glass. When the objective lens is lowered into the oil, more light is
transmitted to the lens.
Because of the increased resolving power, most bacteriological microscopy uses the oil
immersion lens. It is important to be aware that the working distance, the distance between the
specimen and the objective lens decreases with higher magnification. The 100X lens has an
average working distance of approximately 0.1 mm. Care must be taken not to break the slide or
the coverslip when viewing at this magnification.
Become familiar with the brightfield microscope. Use the low and high power dry lenses and the
oil immersion on each slide. Quality brightfield instruments are parfocal and parcentric. This
means that the specimen remains in focus and at the center of the viewing field when objective
lenses are rotated. You should therefore not have to use the coarse adjustment at higher
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Coarse and fine adjustment knobs
Parts of the Microscope
Ocular Lens/ eyepiece
Revolving Nosepiece
Objective lenses
Mechanical Stage Controls
Slide Holder
Stage
Coarse Adjustment knob
Fine Adjustment knob
Condenser
Diaphragm Lever
Light
Light Intensity Adjustment
Arm
Base
Function
Magnifies the image. In BIOL 302L, ocular lens is 10X
Rotates objective lenses into viewing position
Magnifies the image
4X: Scanning
10X: Lower Power
40X: High Dry
100X: Oil Immersion
Moves slide holder/ specimen to view different regions
of the specimen
Holds slide in place and adjust with stage controls
Holds slide and opening to allow light pass to slide
Brings the specimen into focus with the 4X and the 10X
objective
Brings the specimen into focus with the 40X and 100X
objective
Focuses the light on the specimen
Controls the amount of light passing through the
condenser
Illuminates the specimen
Controls the light intensity
Supports the head and stage
Supports the microscope
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magnifications. Once you add oil to your slide, do not return to the dry lenses!
Care and Use of Brightfield Microscopes
1. Always carry the microscope upright and with two hands - one hand should grip the arm
and the other hand should be placed firmly underneath the base. Be careful not to bump
or drop the instrument. Make sure that your lab bench is clear before you bring the
microscope.
2. Use only optically safe lens paper to clean the objective, ocular lenses and the
condenser before and after each lab period. If you should find a microscope that has not
been cleaned, please notify your instructor. Do not use any solvents or soap without the
permission of your instructor.
3. To view a sample, use the objectives sequentially (without skipping) from lowest power
to highest power. First, with the 4X objective, get the sample into focus using the coarse
focus then fine focus; use the stage controls to center cells in the viewing field. Then
switch to the 10X objective; only a slight adjustment with the fine focus should be
needed since the lenses are parfocal. The coarse focus should only be used with the 4X
objective, not when using any of the higher-powered objectives. Then, switch to the 40X
objective, again using the fine focus and centering the sample. Finally, revolve the
objectives so that the slide is between the 40X and 100X objectives. Add a drop of
immersion oil to the slide, and CAREFULLY revolve the 100X objective directly into the
oil drop on the slide. Once oil is on the slide, DO NOT go back to the 40X objective
again. (Why not??) As you move forward from lower-powered to higher-powered
objectives, it’s important to get the sample centered and well-focused, since you
shouldn’t go backward to the 40X once you have used the 100X objective.
4. Use lens paper to clean immersion oil from the 100X objective. Make sure that the oil
does NOT get on the dry objectives (4X, 10X, and 40X).
5. Before returning the microscope to its designated space in the storage cabinet:
a. remove any slide
b. clean the optical system
c. rotate the lowest power objective into position
d. center the mechanical stage
e. wrap the light cord around the base, and cover.
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Other types of microscopes
Fluorescence Microscope: This variation of the brightfield microscope uses a mercury arc vapor
lamp as a source of ultraviolet light. A fluorophore, or a chemical compound with known
fluorescent activity, can be attached to a sample for visualization. When ultraviolet light hits the
fluorophore, the electrons in the molecule become “excited.” When they return to their natural
energy state, they release energy in the form of light, which we can then visualize in the
fluorescence microscope. A filter is between the objective and ocular lenses, which allows only
the fluorescence emitted from the sample to reach the eyepiece. A special condenser is used so
that the background field is dark, thereby resulting in a high contrast fluorescent image. The
color that we visualize after the electrons emit light depends on how much energy the electrons
absorb, which depends on their location in the valence shell. It can be predicted using a graph
called an absorption and emissions profile, which plots relative intensity of fluoresced light
against wavelength. The difference between the absorption peak and the emissions peak is called
the Stokes Shift. Different wavelengths of light will be visualized as different colors.
Fluorescent microscopes are often used in laboratories to visualize localization of certain
compounds within cells, since fluorophores with specificity for certain molecules can be
designed.
(Wikipedia.com)
Confocal Microscope: Confocal Microscopy is similar to fluorescence microscopy but uses
point illumination. By illuminating only the point of focus, unwanted scattered light is reduced.
In addition, light from the specimen passes through a pinhole aperture that blocks light not
coming from the focal point. Reducing stray light from sources not at the focal point improves
resolution. This technique also allows 3D images of the sample to be built from viewing thin
sections, which is useful for looking at thick samples and preserving spatial relationships.
Phase Contrast Microscope: Living cells can best be viewed with a phase-contrast microscope.
The optics of the microscope translate differences between the refractive indices of cellular
components and the surrounding medium into differences in light intensity. The phase contrast
microscope is very useful for looking at small cells at lower magnification (40X).
Darkfield Microscope: In this microscope, the condenser is designed to prevent light from
15
passing to the objective unless it strikes the specimen. Thus the background is dark. The high
contrast between background and the specimen makes it possible to view living unstained
specimens.
Electron Microscopes: Both the transmission electron microscope (TEM) and the scanning
electron microscope (SEM) contain magnetic lenses that focus a beam of electrons on the
specimen. Electrons used in this fashion generate a wavelength that may be 100,000 times
shorter than that of visible light. As a result, electron microscopes have resolving powers as
much as 400 times that of light microscopes and 200,000 times that of the human eye. Electron
microscopy gave scientists their first look at the world of viruses.
The TEM bombards a thin specimen with electrons. Depending on their composition, the
components of the specimen transmit, absorb or deflect the electrons. The image produced on a
photographic plate is a visual translation of this interaction of electrons with the specimen. The
transmission electron microscope exposes many secrets of subcellular structure. Samples are
usually fixed in a resin and then very thin slices are observed allowing us to see a cross section
of a cell.
The SEM is quite different from the TEM. It is designed to generate dramatic images of the
surface that provide information about depth and surface structure. In this microscope an electron
beam is moved back and forth over the surface of a metal-coated specimen causing the emission
of secondary electrons from the specimen. The secondary electrons produce the stunning images
characteristic of scanning electron microscopy.
Microscope Virtual Lab (North Carolina Bionetwork)
As preparation or review of the microscopy lab, work through the microscope virtual lab to learn
about brightfield microscopes and practice your technique:
https://www.ncbionetwork.org/iet/microscope/
The virtual lab will take about 30 minutes to complete.
On the main screen, there are buttons for: Guide, Learn, Explore, Test, Options. We will be
using Guide, Learn, and Explore. (You may use Test to quiz yourself, and Options is for closedcaptions.)
PROCEDURES
1. First, click on Guide. In the Overview Chapter, read all 12 “pages” to gain an overview of the
different parts of the microscope and their uses. Also, read all pages within the Objective
Lenses, Immersion Oil Lens, and Microscope Care chapters. Return to the main screen.
2. Next, click on Learn. This section has 4 main pages which need to be reached sequentially
(you cannot skip ahead): microscope parts, lenses, oil immersion lens, and magnification.
16
•
•
•
In the first 2 pages, you must click on every “?” to change it into a green check mark
before the Next button appears at the bottom of the screen.
The oil immersion lens page dramatically shows what a difference the immersion oil
makes, for the light rays and for image clarity.
The magnification page allows you to mix and match different eyepiece and objective
lenses to achieve different total magnification, and observe their effect on the image
viewed. Notice how the 4X and 10X objectives are best for scanning; when using the
40X and 100X objectives, the viewing field is so small, you may have no idea which part
of the sample you are viewing!
3. Next, click on “Explore.” The virtual lab has many slides for you to practice your microscope
technique. You can adjust the coarse and fine focus and light intensity by using the sliders, and
move to a different viewing field by dragging the sample. The main drawback with this virtual
lab is that the bacterial images using the 100X objective aren’t very good.
If you’d like to try out additional microscope virtual labs (optional), try:
https://virtuallabs.nmsu.edu/micro.php
Post-lab Questions
1. Contrast “magnification” and “resolution.”
2. Which lens is used with immersion oil? Does immersion oil improve magnification or
resolution? Explain.
3. When using the 100X objective to view samples on our BIOL 302L microscopes, what is the
total magnification of the sample? (Hint: see the microscope diagram a few pages before in the
manual)
4. Once there is oil on the slide, why should you not use the 40X objective?
5. When viewing a slide on the brightfield microscope, what is the sequence of steps you should
follow? (This will be a long answer.)
6. List 3 things you have learned from the virtual lab about proper microscope technique.
7. When performing fluorescence microscopy, what kind of light is used and how is a sample
made “visible”?
8. What type of microscopy has the highest resolution? Why?
9. Using Google or PubMed, identify a research paper which shows a fluorescence image of a
bacterial cell (or cells). Paste in the image, the reference in proper ASM format, and a
description of what the image is showing.
17
Kohler Illumination:
18
19
20
21
Step 17. Switch to the 40X Objective
Step 18. Refocus and adjust the iris diaphragm.
Step 19. Now we will move to 100X oil. Make sure you are satisfied with
40X because you cannot move back once oil is added!
a.
b.
c.
d.
Move between the 40X and 100X so there is no objective locked in for use.
Add a drop of oil to the specimen slide.
Click into place the 100X objective.
Refocus and adjust the iris diaphragm.
22
USING MICROPIPETTES
Micropipettes are used to measure and dispense
microliter volumes. Pipetman is a common
micropipette brand. There are 3 sizes of Pipetman,
p20, p200, and p1000. The sizes indicate the amount
of liquid the Pipetman is capable of dispensing.
The p20 can dispense between 2 uL and 20 uL. The
p200 can dispense between 20 uL and 200 uL. The
p1000 can dispense between 100 uL and 1000 uL.
Never use a micropipette for more or less than its
capacity. This changes the calibration and will make
the rest of your pipetting inaccurate.
The Parts of the Pipetman:
The “Plunger”
This is used for obtaining and
releasing the liquid. You will
notice when you push down
that with limit pressure, the
plunger will stop. If you
apply more pressure, it will
go a little farther and then
stop. DO NOT push the
plunger ALL the way down
UNLESS you are dispensing
the liquid. If you do this
when you collect the liquid
you will collect MORE than
you want.
The Dial
This is used
for setting
the amount
of liquid
that you
want to
pipette.
Conversions:
There are 1,000 uL in 1mL so:
0.1 mL would be __________uL
0.85 mL would be __________uL
0.034 mL would be ___________uL
250 uL would be _____________mL
20 uL would be ____________mL
740 uL would be __________mL
1550 uL would be ____________mL
23
The Ejector
This is used
for ejecting
dirty tips
into the
waste
container.
MICROPIPETTING SOLUTIONS VIRTUAL LAB (LabXchange)
This virtual lab is excellent practice and review for using micropipettes.
https://www.labxchange.org/library/items/lb:LabXchange:4eecf5fe:lx_simulation:1
•
It will take about 10 minutes to complete the simulation.
•
After pressing “Start simulation”, choose “Level 1.”
•
In the simulation, after choosing the P20, click on the cartoon bubble to set the volume.
Post-lab Questions
1. What is the purpose of the “first stop” on the micropipette? When, during the pipetting
process, should you push to the first stop?
2. What is the purpose of the “second stop” on the micropipette? When, during the pipetting
process, should you push to the second stop?
3. If you would like to measure 100 µL, what pipettes could you use or not use? How would the
numerical setting on the dial appear?
24
PURE CULTURE TECHNIQUES
OBJECTIVES: Students should be able to
1. Define pure culture and colony forming unit.
2. Compare the advantages and disadvantages of the spread plate and streak plate
procedures.
3. Isolate an organism in pure culture using a spread plate and streak plate procedure.
4. Determine which technique is appropriate for various situations.
Until 1883, bacteria were grown in the laboratory only in liquid broth or on the surface of fruits
and vegetables. One of the most important discoveries in microbiology was the development in
the laboratory of the German scientist Robert Koch of a solid nutritional medium for isolating
pure cultures, a single organism and its reproduced progeny. Microbes could be spread across
the solid surface and separated from each other. When these individual organisms reproduced,
they formed colonies that represent the progeny of a single bacterium (colony-forming unit or
CFU); each colony is a pure culture. For example, each single colony of E. coli on a TSB plate
contains 10,000,000 to 100,000,000 cells that were all descended from one mother cell, so each
cell within a colony is essentially genetically identical to the other cells in the same colony.
The first widely used solidifying agent was gelatin, which is broken down by some bacteria and
melts at temperatures above 30oC. (In the summer it may be impossible to work with gelatin in a
room without air-conditioning.) Because of this limitation, agar, a more stable algal product,
replaced gelatin.
Only with pure cultures can the properties of individual types of microorganisms be examined
and understood. For example, in the 1890’s Koch’s postulates (which built the foundation of
medical microbiology) required the use of pure cultures to demonstrate that certain bacteria
caused a disease.
Bacterial cells are commonly separated from each other by using a quadrant streak plate or
spread plate. In the streak plate, cells are dragged farther and farther apart from each other on the
surface of the plate so, at some point in the streak, single cells and their offspring can form a
distinct colony. Because the streak plate involves a kind of dilution on the surface of the plate
via repetitive streaking with a sterile loop, it can be performed with very dense cultures. Spread
plates need to be performed with cultures that are diluted enough to give a countable number of
colonies. Remember to use aseptic technique to prevent contamination from the environment.
MATERIALS
•
•
TSA or other appropriate agar medium
Petri dishes
25
•
•
•
•
•
•
Glass spreading rod
95% Alcohol
Micropipettors
Inoculating Loop
Cultures in broth for spread plates at concentrations of ~103 org/ml
Cultures at a concentration of 108 org/ml for quadrant streak
PROCEDURES
SPREAD PLATE
1. Pour a plate of TSA and allow to solidify.
2. Suspend the organisms in the broth culture. Remove the cap
from the culture tube and flame the mouth of the tube. Do
not contaminate the cap during this procedure. Remove 100
µl of organisms. Flame the mouth again and cover the tube.
3. Dispense the suspension in the center of the plate.
4. Dip the bent glass rod in the alcohol and shake off excess
liquid. Keep the alcohol container away from the flames.
5. Carefully flame the rod. When all the alcohol has burned,
allow to cool for a few seconds. You may be sure the rod is
cooled by placing it on the agar surface at the edge of the
plate.
6. Use the rod to spread the organisms over the surface of the plate while rotating the plate
on the desktop.
7. Return the glass spreader to the alcohol and flame to sterilize.
8. Incubate the plate for 48 hours in an inverted position. Look for isolated colonies.
QUADRANT STREAK PLATE
1. Pour a plate of TSA and allow to solidify. Return the TSA to the water bath.
2. Suspend the organisms in the broth culture or use directly from a slant.
3. Flame the loop and wire until it is red hot. Remove the cap from the culture tube and
flame the mouth of the tube. Do not contaminate the cap or the loop during this
procedure. Remove a loopful of organisms. Flame the mouth again and replace the cap on
the tube.
26
4. Spread the organism over a small region on the edge of the plate as shown on the left in
the diagram below.
5. Flame the loop and let it cool for a few seconds. Make sure the liquid from the streak has
completely dried (absorbed by the plate) before proceeding with the next streak.
6. Streak from the end of region 1 across the edge of the plate, forming region 2.
7. Repeat step 5.
8. Streak from the end of region 2 across a quarter of the plate, forming region 3.
9. Repeat step 5.
10. Streak from region 3 across the remaining portion of the plate, forming region 4.
11. Flame the loop before setting it down.
12. Incubate the plate for 48 hours in an inverted position. Look for isolated colonies.
Post-lab Questions
1. What are the advantages and disadvantages of the spread plate vs. streak plate technique?
27
2. You expose an agar plate to the environment and get good growth. A number of colonies
overlap and you wish to isolate the organisms in pure culture. Which technique would
you use? Explain.
3. When performing a streak plate, why is it important to allow each streak to dry before
proceeding with the next streak?
4. Suppose your streak plate has single colonies in the third quadrant and no growth at all in
the fourth quadrant. Considering the purpose of a streak plate, do you think that this is a
successful streak plate or not? Justify your answer.
5. How might you prepare a spread plate from organisms growing on a slant?
6. You begin with an overnight culture of approximately 108 organisms/ml and need a
concentration of 103 bacteria/ml in order to get a countable spread plate. Describe how
you would dilute your sample.
7. Before the development of solid media by Robert Koch, why was it impossible for a
scientist to be certain that he/she had obtained a pure culture?
8. Are there any other methods to isolate single colonies in agar? Look in the literature and
provide the reference.
28
PRINCIPLES OF STAINING, BACTERIAL MORPHOLOGY,
SMEAR PREPARATION, AND SIMPLE STAINS
OBJECTIVES: Students should be able to
1. Describe the principles of staining and differentiate between
a. specific and nonspecific stains
b. acid and basic stains
2. Explain the value of stains to microbiology
3. Prepare a smear from bacteria grown in liquid or solid media
4. Perform and interpret a simple stain
Even with the microscope, bacteria are difficult to see unless they are treated in a way that
increases contrast between the organisms and their background. The most common method to
increase contrast is to stain part or all of the microbe.
There are several staining methods that are used routinely with bacteria. These methods may be
classified as 1) nonspecific and 2) specific. Nonspecific stains will react with all microbes in an
identical fashion. They are useful solely for increasing contrast so that morphology, size and
arrangement of organisms can be determined. Specific stains give varying results depending on
the organism being treated. These results are often helpful in identifying the microbe.
Bacterial Morphology
Bacterial morphology generally falls into four main categories: rod-shaped (bacillus; plural
bacilli), spherical (coccus; plural cocci), curved (vibrio), and spiral (spirillum or spirochete).
Examples of specific morphologies are shown on the next page.
How Stains Work
Stains are chemicals containing chromophores, groups that impart color. Their specificity is
determined by their chemical structure. For example, a basic stain is a stain that is cationic
(positively charged) and will therefore react with material that is negatively charged. The
surface of bacteria at neutral pH is somewhat negatively charged and will therefore attract basic
stains. Some examples of basic stains are crystal violet, safranin, basic fuchsin and methylene
blue. In contrast, acidic stains have negatively charged chromophores and are repelled by the
bacterial surface. They stain the background and leave the microbe transparent. Nigrosine and
Congo red are examples of acidic stains.
29
Preparation of bacteria for staining
One or more stains can be used during a staining procedure. When a single stain is used, the
procedure is known as a simple stain. The stain interacts with some portion of the bacterium
determined by the chemistry of the stain and the microbe. This procedure requires preparation of
a smear - a thin layer of the specimen immobilized on a slide before staining. Immobilizing the
sample is important so that excess stain that is not bound to the sample can be washed away.
Air drying helps to immobilize a sample onto a slide, but for best adherence, the sample should
be fixed. Fixation not only improves attachment of the sample to the slide, it also preserves cell
structures and inactivates enzymes that might damage cell morphology. Fixation can be
accomplished using heat (passing the slide through a flame) or chemical fixatives. Heat fixation
is simple and quick, but chemical fixation may be preferred when observing delicate structures
within cells.
You will practice preparing smears and staining organisms from your environmental sample
plate or from pure cultures provided by your TA.
SIMPLE STAIN
MATERIALS
§
§
§
§
§
Clean microscope slides
Methylene blue or other basic dye
Wash bottle
Bibulous paper
Cultures (choose 2 of the following)
§ organisms from your Isolation of Bacteria from the Environment plate
§ Corynebacterium xerosis (slant)
30
§
§
Staphylococcus epidermidis (slant)
Klebsiella pneumoniae or Escherichia coli (slant)
PROCEDURE
1. Prepare two smears (1 smear each with 2 different bacteria), following the instructions
below:
§
Preparing a smear from bacteria grown in liquid medium
Spread one loopful (or 10 uL) of the liquid over a large area of a clean slide. (If a
slide is not clean, the water will not spread out into a thin layer but will form beads.
It is recommended that new precleaned slides be used. Used slides may be cleaned
by scrubbing with a cleanser.) Allow the sample to air dry.
Heat fix your smear to the slide. This attachment process is accomplished by passing
the slide over the flame of a Bunsen burner two or three times. Because the slide may
become hot, it is advisable to use a slide holder. Be careful, however, not to
incinerate the organisms on the slide. Proper fixation does not require extremely
prolonged exposures to heat.
§
Preparing a smear from bacteria grown on solid medium
Place one loopful (or 10 uL) of water on the slide.
Using an inoculating needle, mix a very small quantity of the culture with the water
and spread over the slide. It is critical that microbes are separated from each other
during this step.
Air drying and heat fixation are performed as described above.
2.
Cover with methylene blue for one minute. Do not touch the dropper to the slide. The
amount of stain used should be sufficient to cover the smear without flowing off the
slide. To avoid staining your hands, wear gloves or use a slide holder.
3.
Tilt the slide and gently wash the stain off using the water bottle. Be sure that the stain
runs off into the sink or into a stain pan.
4.
Shake excess water off the slide and blot it gently using bibulous paper – if blotting is
too vigorous, “cracks” or “scratches” will be visible when viewing the sample under
the microscope. If you have the time, you may allow the sample to air dry.
5.
Use Kohler Illumination and examine using the 100X objective lens.
6.
For each slide, diagram and describe the shape and arrangement of the organisms in a
typical microscopic field. Indicate how your technique may be improved in the future.
31
Post-lab Questions
1. What is the purpose of fixation?
2. Would you expect all bacteria to react identically to each other when treated with a
specific chemical dye? Explain your answer.
3. What information can be obtained from a simple stain?
32
WINOGRADSKY COLUMN
OBJECTIVES: Students should be able to
1. Culture and visualize a diverse self-contained microbial ecosystem.
2. Describe how the different metabolic capabilities of the microbes can be inferred by
coloration of the layers, due to pigmentation of the microbes and mineral formation.
The Winogradsky column, first used by Sergei Winogradsky in the 1880s to study a complex
microbial ecosystem, is a small environment where we can observe a diversity of microbes that
thrive under a gradient of oxygen, sulfide, and light.
from http://jan.ucc.nau.edu/~doetqp-p/courses/env440/env440_2/lectures/lec23/lec23.html.
Retrieved March, 2011.
MATERIALS YOU WILL NEED TO BRING (per group):
• A narrow clear plastic bottle with the cap (approximately 7” high and 2” wide -water
bottles or bottles with a wide mouth work great)
• Mud/sand from a forest, garden, lake, marsh, ocean; you will need enough to fill your
bottle approximately 2/3 full
33
•
•
•
Water from the same location as your mud – enough to fill the bottle
A carbon source – finely shredded grass, leaves, newspaper, corn starch, or sawdust, etc.
A sulfur source – a hard-boiled egg yolk, cheese, or calcium or magnesium sulfate
(provided)
MATERIALS PROVIDED:
• Wooden dowel
• Beaker to mix mud in
• Calcium or magnesium sulfate
• Sodium bicarbonate
PROCEDURE:
1. Break up the soil if necessary. You do not want any big chunks. Remove them if
necessary. Make a slurry with the water to the consistency of a milkshake. Add in about
1-2% (v/v) calcium sulfate. Mix well. You may also add some sodium bicarbonate
(1%), but it is not necessary.
2. Add the material to the jar, tapping it down with the wooden dowel as it is added to about
2/3 full. Avoid air pockets.
3. Add additional water if necessary for 2-3 cm of water over the mud.
4. Close the jar LOOSELY – Do not seal cap!
5. Incubate on the windowsill in the light or under grow lights.
6. Over time, describe the changes in the bottle. What types of microbes can you identify?
7. On the “Winogradsky Column results” day, observe the microbes from at least 2 different
layers (the top layers are easiest to reach and have the least odor) under the microscope
using the 40X and 100X objectives. Record your observations in your lab notebook.
8. At the conclusion of the semester, feel free to continue to enjoy your Winogradsky
column in your own home. If you do not want to take it with you, please dispose of the
column in the trash. Do not put the mud down the drain.
Check out: http://beautyinscience.com/Winogradskyfinal.jpg
Post-lab Questions
1. In many microbial ecosystems, including a Winogradsky column, the metabolic
requirements of one group of organisms can be provided by the product of another group
of organisms. Provide an example.
2. Both cyanobacteria and purple sulfur bacteria perform photosynthesis. Why do you find
cyanobacteria on the surface of the soil and purple sulfur bacteria deeper in the column?
3. Describe the morphology of the microbes in several different colored layers.
4. Draw the graph of catalase activity vs. depth in the column that you would expect to see.
Explain your answer.
34
CAPSULE STAIN
When a stain, such as an acid dye, cannot penetrate the outer layers of a microbe, the cell will
appear transparent on a colored background. This stain is called a negative or background
stain and is performed by mixing the dye with a suspension of bacteria on a slide and spreading
the mixture into a thin layer for viewing.
The capsule is a structure surrounding the cell envelope that certain bacteria can produce. The
ability to form a capsule is genetically and environmentally controlled. Only those microbes
with the genes for capsule production have the potential to manufacture this polysaccharide (or
polypeptide) surface layer. Special nutrients or other growth factors often are necessary for the
genes to be expressed. The role of the capsule is primarily for protection of the bacteria. For
example, the capsule binds to water and protects against dehydration. Many capsules repel white
blood cells and thus allow pathogenic invading bacteria to elude one of the primary host
defenses.
Capsules are not readily stained and therefore are visualized by negative stain techniques. The
organisms are prepared as a smear in the presence of an acid dye and allowed to air dry because
heat will cause the capsule to shrink. Our procedure will combine an acidic dye, Congo Red
that stains the background, and a basic dye, Maneval’s stain that stains the bacterial cell. The
capsule appears as a colorless layer between the bacterium and the background.
MATERIALS
•
•
•
•
Congo Red
Maneval’s stain
Microscope slides
Cultures (controls)
§ Positive: Klebsiella pneumoniae (slant)
§ Negative: Corynebacterium xerosis (slant)
Capsule stain with Klebsiella pneumoniae (D. Dinh)
35
PROCEDURES
ROCEDURES
PROCEDURES
PROCEDURES
1) Begin with a drop of Congo Red stain at one end of the slide:
PROCEDURES
2) Aseptically add your bacteria and stir well with a loop to remove clumps.
3) Take a second clean slide and place it on the surface of the first slide and make a smear,
spreading the bacteria and the stain:
4) Air dry. DO NOT HEAT FIX:
5) Add Maneval’s Stain, let sit for 1 minute and rinse with water:
6) Gently blot slide with bibulous paper or paper towels:
7) Using Kohler Illumination and the 100X objective lens, view capsule stain.
!
38
38
!
38
38
36
GRAM STAIN
OBJECTIVES: Students should be able to
1. Explain the biological basis of the Gram stain reaction
2. Describe its value to microbiology as well as its limitations
3. Perform a Gram stain on a gram positive and Gram negative bacterium
4. Perform a Gram stain from a broth culture and agar culture
The
most used
stain in
bacteriology
5. interpret
a Gram
stain
reaction is the Gram stain developed in 1884 by Hans Christian
Gram. This procedure is a differential stain method meaning that this stain can be used to
distinguish between different groups of bacteria. The Gram stain categorizes bacteria into two
groups based on their reaction to the stain, mainly due to differences in the thickness of the cell
wall. The size, shape and arrangement of the organisms can also be determined from a stained
specimen.
The four steps of the Gram stain can be summarized as follows:
1. Primary stain - cover the smear with Gram’s crystal violet for one minute. All
bacteria will take up this dye and appear violet. Rinse off the excess dye with water.
2. Mordant - Gram's iodine is added to interact for one minute with the crystal violet.
This complex will be difficult to remove from certain bacteria during the
next (decolorization) step. Excess iodine is rinsed off with water.
2. Decolorization – Gram’s alcohol (95% ethanol) is briefly (10-20 seconds) applied to
the smear followed by a water rinse.
3. Counterstain - Safranin is added for 20 seconds to dye any decolorized cells, and
excess safranin is rinsed off with water. The safranin will not change the color of the
cells that retain the crystal violet.
GRAM STAIN REACTIONS
STEP
TIME
GRAM +
GRAM -
Gram’s crystal violet
1 min
violet
violet
Gram's iodine
1 min
violet
violet
Gram’s alcohol (95% ethanol)
20 sec
violet
colorless
Safranin
20 sec
violet
pink
37
Gram positive bacteria will appear violet at the end of the procedure, since the thick cell wall
allows the bacteria to retain the crystal violet-iodine (CV-I) complex during the decolorization
step. In contrast, since the thin cell wall of Gram negative bacteria allows the CV-I complex to
be washed out during the decolorization step, these cells will appear pink from the counterstain
at the end of the procedure.
Gram positive and Gram negative cells have very different cell envelope structures (see your
textbook) and thus different characteristics. In clinical laboratories this information can quickly
help in selecting a drug to fight an infecting organism. For example, penicillin is rarely effective
against gram negative bacteria, but often is the drug of choice against gram positive bacteria.
(Penicillin must penetrate the bacterial cell wall in order to interfere with cell wall synthesis.
The Gram negative outer membrane usually prevents entry of penicillin.)
Limitations of the Gram Stain
•
A fresh culture should be used because old Gram positive bacteria may decolorize and
stain with safranin.
•
Some organisms are Gram variable, which means that some isolates appear Gram
positive, some appear Gram negative and some have a mixture of cells that appear Gram
positive and Gram negative.
MATERIALS
•
•
•
Gram stain reagents
Environmental isolate
Cultures (on slants)
§ Gram Negative Control: Escherichia coli
§ Gram Positive Control: Bacillus megaterium and/or Staphylococcus epidermidis
PROCEDURE
1. Prepare smears of each organism. Air dry and heat fix.
38
2. Perform the gram stain as described above and shown in the accompanying table. View
your slides using Kohler Illumination and the 100X objective.
3. Diagram the appearance and arrangement of each organism and indicate the gram
reaction in your notebook.
Post-lab Questions
1. List four cell structure differences between Gram positive and Gram negative bacteria.
2. Why could it be beneficial to perform a Gram stain on a mixed culture?
39
WORKING WITH DILUTIONS
OBJECTIVES: Students should be able to
1. Design a dilution protocol to count viable bacteria
2. Calculate the concentration of a culture from the viable count and the
dilution scheme.
3. Accurately transfer liquids using a pipette
Under optimal growth conditions, bacteria can reproduce until they reach a population density of
approximately 109 (one billion)/ml. Thus, it becomes necessary to dilute them in order to isolate
single organisms, estimate their numbers, or prepare smaller populations for analyses. When
bacteria are suspended in liquid, they are diluted by mixing a measured volume of the culture
with measured volumes of sterile water, buffer or liquid media.
DILUTING: Dilutions are generally performed in multiples of two or ten. This makes it easier
to perform and to calculate concentrations. For example, a 1 to 10 dilution (written 1:10) is 1
volume of bacteria into a total of 10 volumes of liquid. This means that 1 volume of bacteria is
added to 9 volumes of diluting liquid (making a total of 10 volumes).
How would you dilute a liquid culture containing 100,000 organisms/ml to make one containing
100 organisms/ml? You could take 1 ml of the culture and place it into 999 ml of water, but that
would be a cumbersome volume to work with. For reasons of practicality and accuracy,
dilutions of large number of bacteria utilize a serial dilution, a series of smaller dilutions rather
than a single large dilution. The final dilution is calculated by multiplying each of the smaller
dilutions. Thus, if 1 ml of culture is diluted into 9 ml of water (1:10) and 1 ml of this diluted
material is added to another 9 ml of water (1:10), the final dilution will be 1:10 x 1:10 or 1:100.
A third 1:10 dilution would give a final dilution of 1:1000.
To determine the actual number of organisms in each tube, a 0.1 ml sample can be removed and
plated. The ideal number of colonies to count is in the range of 30 to 300 colonies per plate. If
there are more than 300 colonies on a plate, in this course we will call that “too numerous to
count” (TNTC).
CALCULATIONS
Practice your understanding of dilutions with the following problems.
1. You perform a serial dilution.
• First dilution: pipette 1 ml of culture into 9 ml of water.
• Second dilution: pipette 1 ml of first dilution into another 9 ml of water.
• Third dilution: pipette 1ml of the second dilution into 99 ml of water.
40
•
Diagram this procedure. What is the total dilution?
2. You perform a serial dilution starting with a culture containing 1,000,000 organisms/ml.
• First dilution: pipette 1 ml of culture into 99 ml of water.
• Second dilution: pipette 1 ml of first dilution into another 99 ml of water.
• Third dilution: pipette 1ml of the second dilution into 9 ml of water.
• What is the total dilution at each step? What is the number of organisms/ml at each
step?
3. In the previous question, you prepare spread plates from the second dilution by pipetting
either 0.1 ml or 0.3 ml samples. How many organisms would you expect on these plates?
4. You have a culture containing 109 organisms/ml and wish to dilute it to 102 (100)
organisms/ml. Design a serial dilution. (Note: total volume (x+y) should not exceed 50
mL)
5. You have performed the following serial dilutions: 1:100, 1:100, 1:10. Diagram this
dilution scheme and include the overall dilution in each tube. From each dilution, you
then prepare pour plates using either 0.1 ml or 1 ml samples. After incubation you count
the plates and get the following results:
Final Dilution
1:100
1:10,000
1:100,000
Colonies – 0.1 ml
TNTC
63
7
Colonies – 1 ml
TNTC
TNTC
59
What was the number of organisms/ml in the initial culture?
PROCEDURES
1. You have been given a culture that has 108 cells/ml. As a group, given that your diluent
volume in each tube is 9.9 or 9 ml, write out a dilution series so that you have tubes with
106, 104, 103 and 102 cells/ml.
2. Prepare the dilution series you just calculated. On separate plates, plate 0.1 ml from each
of the tubes you have prepared with 104, 103 and 102 cells/ml.
3. Predict how many colonies you expect to see on each plate.
4. Next lab period, count the colonies on your plates. Compare the actual numbers of
colonies with the expected numbers. Based on your colony numbers, what is the original
CFU/ml? In your notebook, explain why the actual and expected numbers may differ.
41
ACID FAST STAIN
OBJECTIVES: Students should be able to
1.
2.
3.
4.
Explain the biological basis of the acid fast stain.
Describe its value to microbiology and its limitations.
Perform and interpret an acid fast stain.
Compare the Gram stain to the acid fast stain.
The acid-fast stain is a differential stain that clinical laboratories will often perform to
distinguish mycobacteria from other bacteria. These organisms include the etiological agents of
tuberculosis, leprosy and some pneumonia associated with AIDS. Bacteria in the genus
Mycobacterium have mycolic acids, special lipid-like material attached to their cell walls, that
renders them impermeable to most aqueous stains.
The Kinyoun acid-fast staining method uses high concentrations of the primary dye
carbolfuchsin (a red dye with 5% phenol). Once stained, acid-fast bacteria retain the
carbolfuchsin after a decolorization step with acid-alcohol. Most other bacteria and tissue cells
would be decolorized by acid-alcohol and can be counterstained with methylene blue. Thus the
acid-fast organisms will be red on a background of blue non acid-fast cells.
Carbolfuchsin/phenol for 3
minutes and rinse
Decolorization with 3% HCl
in 95% alcohol and rinse
Counterstain with methylene
blue for 2-3 mins and rinse
Acid fast organism
Red
Non acid fast organism
Red
Red
colorless
Red
Blue
Photo by P.C. Anguiano
42
MATERIALS
•
•
Acid fast reagents: Kinyoun carbolfuchsin, acid alcohol and methylene blue
Cultures:
• Mycobacterium smegmatis/phlei (slant)
• Staphylococcus epidermidis (broth)
PROCEDURE
1. You will perform the acid-fast stain on a single slide with the two bacteria in
separate smears. Prepare separate smears for M. smegmatis/phlei and S.
epidermidis at two ends of a clean slide. Use a sterile loop to break clumped
material.
2. Let the slide air dry and then heat fix.
3. Cover each smear with Kinyoun carbolfuchsin and stain for 3 minutes. Rinse
with water.
4. Decolorize with acid alcohol for at least 30 seconds and rinse with water. Make
sure to wash out the carbolfuchsin well before proceeding to the next step.
5. Counterstain with methylene blue for 2-3 minutes. Rinse with water, blot dry,
and examine with the microscope using Kohler Illumination and the 100X
objective.
In your notebook, diagram and describe the stain reaction, and the morphology and arrangement
of the organisms in a typical microscopic field.
Post-lab Questions
1. When subjected to the Gram stain, mycobacteria either stain weakly red or don’t stain at
all. Explain why, describing the differences between the cell envelopes of acid-fast, Gram
positive, and Gram negative bacteria.
2. Synthesizing cell walls with mycolic acids requires a great deal of energy. Do you expect
acid-fast bacteria to grow fast or slow compared to other bacteria? Explain.
3. What are the chemical differences between the Gram stain and the acid fast stain?
43
LIVE/DEAD STAIN
OBJECTIVES: Students should be able to
1. Explain the biological basis of the LIVE/DEAD stain reaction.
2. Describe the value of the LIVE/DEAD stain as well as its limitations.
3. Perform and interpret a LIVE/DEAD stain reaction.
4. Propose a hypothesis that can be tested using the LIVE/DEAD stain.
Observations of bacterial cells with brightfield microscopes can yield information about the size,
shape, and arrangement of cells. Additional information can be obtained through the use of
staining. A fluorescence microscope can be used to detect fluorescent probes or dyes that are
applied to a specimen. Two valuable techniques using fluorescence are LIVE/DEAD stains for
bacterial viability and FISH, fluorescent in situ hybridization, for phylogenetic analysis. The
LIVE/DEAD stain is a kit that is produced by Molecular Probes. It contains two fluorescent
nucleic acid stains of different colors. The SYTO 9 green fluorescent nucleic acid stain labels
all cells in a culture whether living or dead. The red fluorescent nucleic acid stain, propidium
iodide, only enters cells with damaged membranes. When both dyes are used, living cells will
stain green and damaged/dead cells will stain red (the SYTO 9 levels are reduced in these cells).
The background is virtually nonfluorescent.
These stains can be used for mixed populations of bacteria from environmental sources or for
pure cultures manipulated in the lab. There are several considerations in interpreting the results
of the LIVE/DEAD stain. The stain reaction relies on the integrity of the cell membrane as a
measure of viability. Some organisms that stain green have intact membranes but may not be
culturable in the media that are available so viability plate counts may not match. Some cells
that stain red and appear to be "dead" may only be damaged and can recover. In all cases, it is
important to prepare the samples correctly. If there is too much background fluorescence it may
be necessary to wash the bacteria to remove growth medium or other contaminating material
before staining.
MATERIALS
•
•
•
•
•
LIVE/DEAD stain kit containing SYTO 9 dye, propidium iodide dye
Nonfluorescent immersion oil
Micropipettes and tips
Fluorescence microscope
Cultures (broth), alive and dead (the “dead” culture was heated to 72°C for 20 min)
• Escherichia coli and/or Staphylococcus aureus
44
PROCEDURE
1. Obtain and label two microcentrifuge tubes “Live” and “Dead.”
2. Pipet 0.2 ml (200 µl) of "Live" bacteria into one tube and pipet 0.2 ml (200 µl) of "Dead"
bacteria culture into the other tube.
3. Using a microcentrifuge, spin down the two bacteria cultures at 13,000rpm for 90
seconds.
4. Carefully pipet the media (liquid) to another microcentrifuge tube, leaving the pelleted
bacterial cells at the bottom of the tube. Replace the media with 200 µl of sterile water
and vortex to resuspend the cells.
5. Micropipet 5 µl of the "Live" culture near one end of a very clean microscope slide. (In
step 6 you will place 5 ul of the “Dead” culture on the other end of the same slide.) Add 5
µl of LIVE/DEAD®BacLight™ to the “Live” droplet. Avoid getting the label on your
hands by wearing gloves since these stains bind to nucleic acids and are therefore
potentially mutagenic. Seal the coverglass with fingernail polish.
6. Repeat step #5 above with the “Dead” culture, placing the drop on the other end of the
SAME slide. It is a good idea to pipet, stain, and seal each specimen onto the slide one at
a time, as described in steps 5 and 6, to prevent cross-contamination.
7. Once the nail polish is dry, label your slide with a Sharpie and transport the sealed
microscope slide in the microscope holder to the microscope room and observe the slide
using a fluorescence microscope with the appropriate filter sets. Leave your gloves in the
lab-do not bring them into the microscope room.
8.
Take 4 pictures of your bacteria (one picture with each filter for the “Live” sample, and
one picture with each filter for the “Dead” sample) and save them to the hard drive.
Download them at a later time (when all groups are finished).
9.
Discard your live/dead slide in the “Contaminated broken Glass” waste because these
chemicals are potentially mutagenic.
Post-lab Questions
1. Describe how the LIVE/DEAD stain distinguishes between living and dead cells.
2. Explain limitations of the LIVE/DEAD stain.
3. What results would you expect if the LIVE/DEAD stain was used on endospores? Explain
your answer.
45
DIRECT COUNTS USING A HEMOCYTOMETER
(Read background information in the section, “Quantification of bacteria”)
The Hemocytometer (counting chamber) is designed to hold a known volume. The chamber
contains a grid that can be viewed in the microscope. The area of the different squares of the
grid also represents a known volume. For example, each large square bounded by the double
lines in the illustration is 1 square mm.
MATERIALS:
•
•
•
•
Hemocytometer
Coverslips
Methylene blue
Cultures (broth)
• Escherichia coli or B. megaterium
PROCEDURES
1. First, pipet up and down several times in the culture tube to resuspend the cells and break
up clumps.
2. Pipette 100 µl of the broth culture (avoiding any clumps at the bottom of the tube) into a
tube containing 100 µl of methylene blue.
3. Use a counting chamber with the coverslip on the chamber. Add the diluted sample to
one of the V-shaped slots with a micropipette and the chamber will fill through capillary
action. You only need a small amount because the entire volume of the grid is less than
0.01 ml.
4. Focus on the grid using low power and then carefully switch to high power. The chamber
is thicker than a typical slide so make sure that you clear the objective. View the squares
using high power (40X objective).
46
5. Count the number of bacteria in five of the large 1/25 mm2 squares (the 4 squares in the
corners and the square in the center). If there are too many organisms per square you may
have to go back and dilute your original sample further (such as 1:20 or 1:100). Aim for
100 cells or less per square. Count organisms on the left and upper lines as part of the
square but not those on the right and lower lines. Divide this total number by 5 to find the
average number of bacteria per large square.
6. Calculate the cell concentration (number of bacteria per ml) in your original sample.
Cell concentration = [(average number of cells per large square) / (4X10-6 ml)] X
(D.F.)
47
MEDIA
OBJECTIVES: Students should be able to
1. Distinguish between defined and complex media
2. Locate information about the components of media and explain the purpose of
each component
3. Describe media used for the isolation of specific bacteria
4. Determine the appropriate sterilization technique for various items
5. Evaluate whether a sterilization procedure/equipment is functioning properly
There are many different media used for culturing bacteria. To be useful for culturing a
particular microbe a medium a) must supply the microbe with its basic requirements for growth,
and 2) lack any inhibitory substances. The basic requirements for growth include water, sources
of carbon and nitrogen, essential elements (minerals), and any compound necessary for growth
(amino acids, vitamins) that the microbe cannot synthesize. In addition, the pH, osmotic
conditions, and physical environment must be able to support growth.
All media can be categorized as defined or complex. Defined media are composed of known
amounts of specific chemical compounds. An example of a defined medium you will be using
later on in this course (for the growth curve and Ames Test labs) is minimal media, which
provides only the nutrients necessary for the growth of a microbe. A common minimal medium
recipe contains only salts, sugars, and minerals such as NH4Cl, glucose, and FeSO4. Complex
media contain nutrient-rich substances such as yeast extract, peptone, or tryptone and therefore
the precise chemical constituents are unknown. Some examples of complex rich media are TSB,
LB, and nutrient broth.
Some media can also be categorized as selective, differential, or enriched. A selective medium
permits growth of one group of organisms while inhibiting growth of some other groups due to
specific substances in the culture medium. A differential medium contains an indicator or dye
that gives a visible color change if a certain biochemical reaction has occurred, which is helpful
for diagnosis. An enriched medium contains special ingredients such as blood to support the
growth of many fastidious bacteria, such as those isolated from body fluids.
Examples of specific media:
Trypticase Soy Agar (TSA) – a complex rich medium. Supports the growth of many microbes;
contains tryptone and enzymatically digested soybean meal.
Mannitol Salt Agar (MSA) – a selective and differential medium. The selectivity is obtained by
48
the high (7.5%) salt concentration that inhibits growth of many groups of bacteria but allows the
growth of Staphylococci or other salt-tolerant bacteria. The mannitol and pH indicator in this
medium help in differentiating pathogenic from nonpathogenic Staphylococci. The
uninoculated media has a red color. Many pathogenic bacteria can ferment mannitol to form
acid, making a yellow zone in the media, unlike many non-pathogenic strains.
MacConkey Agar (Mac) – a selective and differential medium. Selectivity of this medium is
due to the bile salts and crystal violet that inhibit the growth of gram-positive bacteria but allow
the growth of Enterobacteriaceae and related gram-negative rods. Because lactose is the sole
carbohydrate and a pH indicator is present, lactose-fermenting bacteria produce colonies that are
various shades of red, whereas non-lactose fermenters produce colorless colonies.
Eosin Methylene Blue Agar (EMB) – a selective and differential medium. The eosin and
methylene blue dyes make EMB a selective media, inhibiting the growth of gram-positive
organisms and allowing the growth of gram-negatives. In addition to the two dyes, EMB also
contains lactose, so that lactose fermenters such as E.coli produce dark colonies with a green
metallic sheen and non-lactose fermenters such as S. typhi appear colorless.
Blood Agar – an enriched and differential medium. The addition of citrated blood to trypticase
soy agar makes possible variable hemolysis, which permits differentiation of some species of
bacteria. Three hemolytic patterns can be observed on blood agar.
1. a-hemolysis - greenish to brownish halo around colonies (e.g. Streptococcus gordonii,
Streptococcus pneumoniae).
2. b-hemolysis - complete lysis of blood cells resulting in an area of clearing around
colonies (e.g. Staphylococcus aureus and Streptococcus pyogenes).
3. g-hemolysis - no change in the medium (e.g. Staphylococcus epidermidis and
Staphylococcus saprophyticus).
Hektoen enteric agar (HEK) – a selective and differential medium. Selectivity is due to the
bile salts that inhibit growth of gram positives. Most species metabolize the sugars in the media,
making a yellow or red color due to the pH indicator, but Salmonella and Shigella species
metabolize the amino acids in the media, creating a blue color. H2S-producing Salmonella make
black colonies whereas non-H2S-producing Shigella make green colonies.
The key difference between a broth and solid media (i.e. TSB vs. TSA) is that agar is included
as a solidifying agent in solid media. Agar is a polysaccharide produced by algae. These days
the majority of commonly used media are purchased in dehydrated form. To prepare it in its
usable form, one only has to add water, dissolve the contents, and sterilize. The principles of
sterilization and the currently used methods of sterilization were discussed earlier in the course.
Our media is generally sterilized using an autoclave.
Part 1. Different types of media.
49
MATERIALS
Media
1. H2O (Water Agar)
2. TSA (Trypticase Soy Agar)
3. MSA (Mannitol Salt Agar)
4. Mac (MacConkey Agar)
5. EMB (Eosin Methylene Blue Agar)
6. Blood Agar
Cultures
1. Escherichia coli
2. Proteus vulgaris
3. Staphylococcus aureus
4. Staphylococcus epidermidis
5. Salmonella enteritidis
PROCEDURES
1.
2.
Each group gets 2 of each type of media plate. Use a Sharpie to label three sections
on the bottom of one plate and two sections on the other plate, so that all 5 organisms
can be inoculated on each type of plate.
Incubate until next lab period.
Part 2. Cellulose production.
Among the dazzling repertoire of activities performed naturally by microbes is the secretion of
strands of cellulose by Acetobacter aceti (xylinum). For its size, the bacterium synthesizes this
product at an impressive rate. Every minute it releases cellulose strands that are 2 microns in
length. The cellulose produced by Acetobacter forms a pellicle. At the surface of a liquid
medium, the aerobic organism becomes trapped in the pellicle and assures itself access to
oxygen. The pellicle also reduces the amount of ultraviolet light that strikes the bacteria. This
reduces the frequency of mutations and thus increases the rate of survival.
Industrial microbiologists are interested in Acetobacter aceti ssp. xylinum because of its potential
as an alternative source of cellulose. Hopefully, it may someday be possible to harvest enough
of this bacterial metabolite to clothe large populations.
The lab instructor will inoculate TSB and Cellulose Supporting Medium (CSM) with
Acetobacter aceti ssp. xylinum. The flask will be incubated at room temperature.
CSM (per liter): 5 gm yeast extract; 3 gm peptone; 25 gm mannitol. Is this a defined or
complex medium?
RESULTS
50
Part 1. Record your observations of growth on the plates in your notebook and answer the
following questions.
• How do these media differ from each other?
• For what practical purposes might you use each of these media?
• Which components give each medium its characteristic properties? For more detail, you
can refer to the Difco Manual, a reference book on dehydrated culture media and to the
Hardy Diagnostic Manual available online. Use these resources whenever you have
questions about the composition, preparation and use of commercial media.
Part 2. For Cellulose production, observe the cultures over the course of several weeks and
record your observations. Do not open the flasks until there is abundant pellicle formation.
Post-lab Questions
For each of the following, show all math.
1. You wish to have sufficient TSA to make 10 pour plates. Each plate holds approximately
20 ml of agar. The instructions on the container are that 40 grams are to be used to make
a liter. How much dehydrated medium do you need?
2. You wish to make 400 ml TSA with a final concentration of 2% NaCl so that it mimics
seawater. TSA contains 0.5% NaCl. How much NaCl do you have to add?
3. Your experimental protocol requires that you make a medium with the following
composition. How much of each component would you use if you were making 500 ml?
§ Tryptone 0.25%
§ Yeast extract 0.05%
§ NaCl 2%
§ NaH2PO4 0.06%
§ MgSO4 0.5%
§ FeSO4(7H2O) 0.002%
§ CaCl2 0.01%
4. How does an enriched medium compare to an enrichment medium?
5. Design an experiment to test the UV protection provided by the cellulose pellicle of
Acetobacter aceti (xylinum).
51
ANAEROBIC CULTURE TECHNIQUES
OBJECTIVES: Students should be able to
1. Distinguish between the categories of bacteria based on responses to oxygen
2. Explain the biological basis for each category
3. Culture aerobes, facultative anaerobes, microaerophiles, and aerotolerant anaerobes
4. Describe how the anaerobic growth chamber works
Bacteria are often categorized according to their growth responses to atmospheric oxygen. This
may vary from species that can grow only in the presence of oxygen to those that can grow only
in the absence of oxygen. The strict or obligate aerobes must have oxygen to grow. The strict
or obligate anaerobes are killed by even trace amounts of oxygen because they lack catalase
and superoxide dismutase, and thus cannot protect themselves from the toxic byproducts
formed during metabolism with oxygen. Strict anaerobes ferment in the absence of oxygen.
Facultative anaerobes can grow in either the presence or absence of oxygen. If oxygen is
available, facultative anaerobes respire with oxygen; if oxygen is not available, they ferment or
use anaerobic respiration (depending on the type of bacteria; only certain bacteria have the
ability to perform anaerobic respiration). Aerotolerant anaerobes are indifferent to the presence
of oxygen, since they ferment in the presence or absence of oxygen. Microaerophiles prefer to
grow in low oxygen concentrations, but can grow without oxygen. Since microaerophiles have
low amounts of catalase and superoxide dismutase, high levels of oxygen inhibit enzymes critical
for growth and are toxic to microaerophiles.
Fluid thioglycollate broth is a reducing medium, that is, it contains compounds that react with
molecular oxygen keeping the free levels low. It also contains the indicator dye resazurin,
which turns pink in the presence of oxygen. Since oxygen is present at the surface of the
medium, the upper layer is usually pink whereas the dye is colorless in the remainder of the tube.
Agar is also included in this medium to give it a semisolid consistency. Fluid thioglycollate
broth has something for every microbe. Strict aerobes will grow only at the top of the tube, strict
anaerobes only at the bottom, facultative and aerotolerant anaerobes throughout the tube, and
microaerophiles somewhat below the surface.
52
Anaerobic jars such as the GasPak are vessels in which an anaerobic environment is generated
after inoculated media are sealed into the chamber. An anaerobic environment is achieved by
adding water to commercially available gas generator envelopes that are placed in the jar just
prior to sealing. Chemicals in the envelope produce hydrogen gas and carbon dioxide. In the
presence of a palladium catalyst, the hydrogen combines with free oxygen in the chamber to
produce water. The carbon dioxide is required for the growth of certain organisms. A resazurin
indicator strip is usually placed in the jar. It turns colorless when the oxygen has been removed.
The candle jar is used to create microaerophilic conditions. It is a large screw-capped container
into which the medium is placed along with a candle. The candle is lit and the jar is sealed. The
candle will burn and reduce the oxygen concentration.
MATERIALS
•
•
•
TSA plates
Fluid Thioglyocollate Broth
Cultures (in broth)
• Clostridium sporogenes (strict anaerobe)
• Escherichia coli (facultative anaerobe)
• Bacillus subtilis (strict aerobe)
• Neisseria sicca (microaerophile)
PROCEDURES
Fluid Thioglycollate:
1. Check to make certain that no more than 20% of the upper portion of the medium is pink.
2. Inoculate the tube to the bottom and gently rotate between the palms of your hands to
disperse the organism. Do not shake or oxygen will be added to the medium.
3. Leave cap loose. Incubate the tubes for 48 hours.
4. Report pattern of growth for each organism.
53
Anaerobic Jar and Candle Jar:
1. Obtain 3 TSA plates and use a Sharpie to label quadrants on the bottom of each plate. On
each plate, streak all four organisms (one in each quadrant).
2. Place one plate in the anaerobic jar. When all plates are in the container, add two gas
packs and an indicator strip. Seal the jar and incubate for 48 hours. The indicator should
be checked to make certain that all oxygen has been removed.
3. Place one plate in the glass candle jar. When all plates are in the jar, place a lit candle in
the jar and close the cap. When the candle has stopped burning, place the jar into an
incubator for 48 hrs
4. Place the third plate in the incubator as an aerobic control.
5. Observe growth.
Post-lab Questions
1. Complete the following chart
Organism
Growth in
Metabolizes
presence of
oxygen
oxygen
Growth in
absence of
oxygen
Detoxifies
byproducts of
oxygen
Aerobe
Facultative anaerobe
Strict anaerobe
Aerotolerant
Microaerophile
2. Compare the results with growth on the aerobic control plate and describe the oxygen
requirements of your organisms.
3. If an organism is a strict anaerobe, it cannot survive in the presence of oxygen even for a
short time. How might you culture these organisms?
4. An aerobe, facultative anaerobe, strict anaerobe, and an aerotolerant anaerobe are
inoculated on a medium in a sealed container with oxygen present. Explain what will
happen.
54
MAINTAINING CULTURES
OBJECTIVES: Students should be able to
1. Distinguish between the roles of a working and stock culture
2. Prepare working and stock cultures
3. Describe how to store working and stock cultures
4. Describe when it is appropriate to create a new working culture from a stock
Cultures may be maintained on media and stored under conditions that inhibit growth. Long
term storage requires other techniques such as lyophilization or storage in liquid nitrogen. In this
course, you will be responsible for maintaining viable cultures of control bacteria and your
environmental unknown. For every organism that you wish to maintain, you must create a stock
and working culture.
The stock culture is an inoculated slant that is incubated under conditions for microbial growth
and then stored at a temperature below the minimum required for growth of the culture. Your
stock cultures should be stored in the refrigerator after its growth on the slant. Stock cultures
are not used unless the working culture dies or becomes contaminated. The stock culture must
be restreaked whenever it is used or at appropriate intervals (approximately every three weeks) to
insure the viability of the organism.
For your group projects you will maintain your own stock cultures in the refrigerator.
The working culture is a slant that is maintained to inoculate media and to make smears for
stains. When needed, a working culture should be generated from the stock culture.
•
Make sure that all cultures are properly labeled and sealed.
•
The name of the organism and other identifying characteristics such as strain type must
be legible and written in permanent marker.
•
The date of inoculation and your initials must also be indicated on the tubes.
•
Write the name of the medium and indicate if it is a stock culture.
•
After sufficient growth store the organism in the appropriate refrigerator. Live cultures
should never be stored in the same refrigerator as toxic volatile chemicals.
55
STERILIZATION
Sterilization: The killing or removal of all living organisms and viruses from an object or area
Heat Sterilization
For liquids (media, buffers)
and solids (tools, supplies,
labware)
How does it work?
Denaturation of proteins
(loss of structure and
function at very high
temperatures), degradation
of nucleic acids, disruption
of membranes
Examples
•The autoclave: Sealed device
that allows the entrance of steam
under pressure. Moist heat
(steam) penetrates objects much
faster than dry heat (oven).
Usual cycle: 15 psi, 121oC, 1520 min.
The high temperature and
pressure kills all
microorganisms – even bacterial
endospores.
Radiation Sterilization
For decontamination of
surfaces (UV) or
plasticware such as petri
dishes (g rays).
Filter Sterilization
For heat-sensitive liquids
Each type of
electromagnetic radiation
(UV, X-rays, g rays,
electrons…) acts through a
specific mechanism that
leads to the death of the
irradiated microorganisms.
The pores of the filter are
too small for the passage of
the microorganisms but
large enough to allow the
passage of the liquid.
56
Not useful for heat-sensitive
liquids.
•UV rays -> DNA damage. The
UV light of a sterile hood is
used to decontaminate the inside
surfaces after use.
•Ionizing radiation (X-rays, g
rays) -> extremely powerful;
DNA, protein, and membrane
damage through ion production.
Standard bacteriological filter
has pores 0.22 µm in diameter,
which will not let bacteria
through. The filters are usually
pre-sterilized by ionizing
radiation.
QUANTIFICATION OF BACTERIA
OBJECTIVES: Students should be able to
1. Perform a standard plate count of a culture in broth
2. Prepare a standard curve of a culture in broth
3. Compare quantitation methods: direct counts, plate counts, filtration, and turbidimetry
4. Estimate the concentration of a broth culture using McFarland standards
Often it is necessary to know the number of bacteria in a specimen, for example, to ensure that
water, milk or other foods are safe to consume. The growth rate or the change in microbial cell
numbers with respect to time is an important characteristic of microbes. Enumerating microbial
populations is also important for evaluating such products as antibiotics, vitamins, and
preservatives. Quantification is also necessary to prepare inoculum for bioassays, tests that
utilize living microbes as the indicator organisms to determine the concentration of a chemical in
solution. These tests are performed by inoculating a known concentration of bacteria into the
solution and measuring growth over time.
Several methods can be used to determine bacterial concentrations. These include direct counts,
plate counts, filtration, and turbidimetric measurements. Bacterial concentrations can also be
estimated using McFarland standards.
Direct counts are usually performed by pipetting a measured volume of fluid onto a special slide
called a counting chamber. The slide is examined under the microscope and the organisms are
counted. Although this is a rapid means of determining concentration, both living and dead
organisms are counted.
Plate counts commonly involve spreading a measured volume onto solid media using the spread
plate technique. If the original sample has a high concentration of bacteria, dilutions are
prepared and plated. The plates are incubated and the number of colony-forming units (CFU)
reflect the viable organisms in the sample. You performed a Plate count during the Working with
Dilutions lab.
Filtration is usually advisable when the number of microbes in a liquid specimen is very low
(<10/ml). A large volume of liquid may be passed through the standard bacteriological filter
(pore size 0.22μm) which traps cells on its surface. The filter paper is placed on a nutrient
medium and incubated and CFUs are counted. Cells on the filter may also be directly counted
after labeling cells with a fluorophore and using fluorescence microscopy.
Turbidimetry is a rapid method of estimating the number of bacteria in solution using a
spectrophotometer. Bacteria absorb light in proportion to their total cell volume (determined by
size and numbers). When microbes increase in number or size in liquid culture, there is an
57
increase in the turbidity (cloudiness) of the culture. Turbidity can be measured as optical
density (absorption of light, usually measured at a wavelength between 520 nm and 700 nm).
For any given microbe, a standard curve can be established relating the number of
organisms/ml (determined by standard plate count) to an optical density measurement
(determined by spectrophotometry). The standard curve describes the relationship between
microbial counts and optical density under defined conditions and can be used to determine
population size measured by optical density readings.
QUANTIFICATION PROCEDURES
DIRECT COUNTS
Microbes are typically counted in a microscope using
• a special slide called a hemocytometer, traditionally used to count blood cells OR
• acridine-orange stained and filtered samples
Hemocytometer
The length, width and height to the coverslip of a hemocytometer are fixed dimensions. Thus
liquid added to the hemocytometer fills a space of known volume. Bacterial cells are counted
within the grids and this number is used to determine the total number of bacteria per ml. The
concentration of the original suspension is determined after adjusting for dilution factors.
Acridine orange stain
Quantitative counts can also be determined by filtering a known sample volume, staining the
cells trapped on the filter with acridine orange, and viewing the filter with an oil immersion lens.
A special filter that does not stain with the dye must be used. The relative area of the
microscopic field to the size of the entire filter can be determined. Several random fields are
counted and the average number of bacteria per field is determined. This number is used to
calculate the total number of bacteria on the entire filter. The concentration per ml of the
original suspension is calculated by dividing by the filtered volume (in mls) and multiplying by
the total dilution factor.
STANDARD PLATE COUNT
A standard plate count is valid if the number of colonies growing on the agar is between 30 and
300. When a suspension contains a higher concentration of bacteria, serial dilutions are prepared
and plated. The concentration of the original culture is determined by considering the number of
colonies, the volume plated, and the dilution factor (for those plates yielding between 30 and 300
colonies).
Generally, an overnight culture of bacteria will be turbid and will contain between 108 and 109
bacteria/ml. It is rare to find samples from nature with higher concentrations. These limits
therefore typically define the dilution series which must be made to evaluate unknown
58
concentrations of bacteria, such as the dilution series and plate count you performed during the
Working with Dilutions lab.
TURBIDIMETRIC MEASUREMENTS
For quantitation by turbidity, a standard curve, a graph of optical density (O.D.) versus
bacteria/ml derived from plate counts, must be available. Once this has been created, the
concentration of the organism can be determined solely by spectrophotometry. A
spectrophotometer directs a beam of light (of a specified wavelength) at the sample and measures
the light transmitted (total light less absorbed light) with a photodetector. The amount of light
passing through the sample is indicated either as optical density (O.D.) or absorbance. The light
is scattered by the cells, so less light is detected after passing through the sample.
Optical density is only proportional to cell concentration within a certain range of cell
concentrations (called the “linear range” of the spectrophotometer). Thus, highly concentrated
cultures must be diluted to give optical density readings that are within the “linear range” where
optical density is proportional to cell concentration; then, the dilution factor is used to calculate
the concentration of the original culture. Similarly, if the cell concentration is too low, the
optical density reading on the spectrophotometer will not accurately reflect the cell
concentration.
McFarland Scale and Standards. The McFarland Scale is a scale numbered from 1 to 10,
which represents specific concentrations of bacteria/ml. It is designed to be used for estimating
concentrations of Gram negative bacteria such as E. coli.
No. Bacteria(x106/ml)
300
600
900
1200
1500
1800
2100
2400
2700
3000
McFarland Scale
1
2
3
4
5
6
7
8
9
10
McFarland Standards are tubes labeled 1 through 10 and filled with suspensions of Barium salts.
Each tube approximates the turbidity of bacterial solutions corresponding to the McFarland Scale
number. Thus tube 7 represents the turbidity of bacteria at a concentration of 2.1 x 109/ml. If
you have a culture and wish to quickly determine its approximate population, its turbidity can be
visually compared to a set of McFarland Standards. If its turbidity falls between tubes 7 and 8,
then the number of bacteria/ml will be between 2.1 and 2.4 billion per ml. The advantage of
these standards are that no incubation time or equipment is needed to estimate bacterial numbers.
59
QUANTIFICATION USING TURBIDIMETRY
MATERIALS
•
•
TSB (Tryptic Soy Broth)
Cultures:
• Escherichia coli (about 108 org/ml)
Preparation of serial dilutions
a. Label spec tubes (near the top of the tubes so the writing doesn’t interfere with the
spectrophotometry reading) for the standard curve - undiluted, TDF 2, TDF 4, TDF 8,
and TDF 16. The unknown culture may not have to be diluted to TDF 16.
b. Pipette 3 ml of TSB into the tubes labeled with TDF. DO NOT ADD TSB TO THE
UNDILUTED TUBE.
c. Pipette 3 ml of the overnight culture into the tubes labeled “undiluted” and “TDF 2”.
d. Carefully mix the TDF 2 tube using a sterile pipette, and transfer 3 ml to the tube labeled
TDF 4.
e. Prepare the TDF 8 and TDF 16 dilutions in a similar manner using a sterile pipette for
careful mixing and transfer. Each tube should contain at least 3 ml; the final dilution tube
will contain 6 ml.
Recording optical density
a. Blank the spectrophotometer using a spec tube containing only TSB in the spec tube
holder.
b. Read diluted material at 595 nm from the most dilute to the most concentrated. Mix to
resuspend particles. Place the spec tube in the spec tube holder. Close the lid and record
the optical density.
c. Repeat using the material in the TDF 8 tube, then the TDF 4 tube, etc.
d. Record your results in your lab notebook.
e. For the standard curve, prepare a graph using a spreadsheet of bacterial concentration vs.
O.D. This graph can be used in later experiments.
Post-lab Questions
1. What are the advantages and disadvantages of the various quantitation methods?
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2. Why must you use a new pipette tip to make each dilution?
3. Explain why a standard curve is valid only for a single microbe in one type of medium.
4. Why must an uninoculated control be used for turbidimetric readings?
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ENDOSPORE STAIN
OBJECTIVES: Students should be able to
1. Define what an endospore is, why it is produced, and what benefits it provides.
2. Perform an endospore stain.
The Schaeffer-Fulton endospore stain is a differential stain to detect endospores in bacterial
cells. Only certain bacterial genera, such as Bacillus and Clostridium, are able to form
endospores. Bacteria form endospores not for reproduction but rather for survival when
starvation conditions are encountered. An endospore is a dormant, metabolically inert form of
the bacteria that is highly resistant to heat, drying, radiation, and chemicals. Although
endospores can survive boiling temperatures, some can be killed by autoclaving, but for others,
glassware must be acid washed prior to autoclaving. Endospores can survive in their inert form
for hundreds of years. Then, given the proper nutrients and growth conditions, endospores can
germinate and resume vegetative growth.
The location of the endospore within the mother cell depends on the species of bacteria. During
sporulation, endospores might be located in the middle of the cell (central), at the end (terminal),
or between the middle and end (subterminal). A mature endospore is released once the mother
cell has lysed.
Endospores are impermeable to most stains, so during the primary staining step with Malachite
green, the smear must be heated to drive the stain into the endospores. Since Malachite green is
water-soluble, the cells decolorize with a water rinse while the endospores retain the stain.
Safranin is used to counterstain the cells.
MATERIALS
Malachite green
Safranin
2-day old slant of Bacillus megaterium or Bacillus subtilis
5-day old slant of B. megaterium or B. subtilis
PROCEDURE
1. Choose either B. megaterium or B. subtilis to work with. For both the 2-day old and a 5-day
old sample of the same species, prepare a smear, air dry, and heat fix.
2. Use the procedure shown on the next page, noting the following: in (a) the paper towel
should be smaller than the slide, and in (c) the paper towel should remain moist with the
62
stain. Whether you are using the steam bath or the flame to heat the slide in (c), do NOT
allow the slide to dry out; add additional malachite green as needed to keep the paper towel
moist. Step (c) will be performed in the hood. Be sure to wear eye protection.
3. Diagram the appearance and arrangement of each organism.
Post-lab Questions
1. How do your two samples look different from one another? Are your results what you
expected?
2. Based on your results, sketch what you would expect if you performed this stain with
bacteria from (a) a 1-day old slant and (b) a 10-day old slant.
3. Suppose you took a loopful of cells from the 5-day old slant and transferred them to a
fresh slant. If you performed the endospore stain with cells that had grown on the fresh
slant for 1 day, what would you expect to observe?
63
64
BACTERIAL IDENTIFICATION VIRTUAL LAB
OBJECTIVES: Students should be able to
1. Explain why the 16S rRNA gene is commonly used to identify bacterial species.
2. Describe how DNA sequencing would be used to identify a bacterial species in a
clinical sample.
3. Understand the purpose of a BLAST search in identifying bacterial species.
For much of this course, you are culturing bacteria and performing biochemical tests to identify
bacterial species based on differences in their metabolism. However, another important method
to identify bacterial species involves DNA sequencing. One major advantage of DNA
sequencing to identify bacterial species is that it can be performed with non-culturable strains,
or strains that cannot be grown in a laboratory because the appropriate growth conditions have
not been discovered yet. It is believed that most microbes on earth are non-culturable.
The gene that is most often sequenced to help identify prokaryotic species is the gene that
encodes the 16S rRNA, the RNA that is a structural component of the small subunit of the
ribosome. The use of the 16S rRNA gene in
prokaryotes (and 18S rRNA gene in
eukaryotes) was pioneered by Carl Woese and
colleagues to demonstrate that the Archaea
represented a domain separate from the
Bacteria. This gene is particularly useful for
identifying Bacteria and Archaea because it is
present in all life and performs the same
function in all organisms. A diagram of the
secondary structure of the 16S rRNA is shown
here. Some regions of the 16S rRNA
sequence are more highly conserved than
others. For example, sequences in the doublestranded regions tend to be more highly
conserved than sequences in the singlestranded regions. Thus, 16S rRNA sequences
from closely-related bacterial species will be
more similar to each other than 16S rRNA
sequences from distantly-related species.
After sequencing the 16S rRNA gene from a
bacterial sample, the sequence is compared against a database containing 16S rRNA gene
sequences from all known species using a BLAST search (Basic Local Alignment Search Tool),
which is a common technique in bioinformatics.
65
This virtual lab demonstrates the procedure that could be used to identify an unknown bacterium
from a patient or from the environment. First, genomic DNA is isolated from a clinical sample.
Second, PCR is used to amplify the 16S rRNA gene from this sample. Third, the DNA is
sequenced. And fourth, the sequence is matched against the NCBI (National Center for
Biotechnology Information) database to identify the bacteria. You will begin the virtual lab
after the DNA from a clinical sample has been sequenced, and you will perform a BLAST search
to match the sequence against a database to identify the bacteria in the sample. (You are
welcome to work through the entire virtual lab from the beginning, starting from "Sample
preparation", but this is not required.)
PROCEDURE
1. Go to http://www.hhmi.org/biointeractive/bacterial-identification-virtual-lab
2. Click "Start virtual lab" (make sure popups are allowed).
3. Click lab picture to enter the lab. Click "Start over" if necessary (i.e. if someone else just
performed the virtual lab on the same computer).
4. At the top, click on the “Intro” tab and read the Introduction. It is helpful to click on extra
links, such as "Learn more about ribosomal RNA".
5. In the left window, click on "6. Sequencing analysis". Read the information here, also reading
the extra information available through the links on this page ("Learn about the science behind
sequence matching" and "learn more about BLAST search results"). Follow the steps in the
“Notebook” window to identify the sample.
6. After you have correctly identified the bacteria in sample A, screenshot the results page that
indicates the correct identification and include it in “Results” in your notebook. Then, click on
the "Samples" tab and choose another sample to identify. Screenshot the results page after you
have correctly identified the second bacteria and include it in “Results” in your notebook.
Post-lab Questions
1. Why must PCR be part of this procedure?
2. How would you ensure that it is the 16S rRNA gene that is amplified via PCR and not some
other sequence from bacterial or human DNA?
3. Why is the 16S rRNA sequence commonly used for identifying bacteria?
4. The Introduction to the virtual lab mentions 16S rRNA and 16S rDNA. What is the difference
between these two? Specifically, where would each be found in a cell?
66
GROWTH CURVE
OBJECTIVES: Students should be able to
1. Describe the phases seen in the growth curve of a batch culture.
2. Predict how different factors influence the shape of a growth curve of a batch culture.
3. Compare the growth curves of batch cultures grown in six different conditions.
4. Determine the generation time for each culture based on the data gained from the growth
curve.
Bacterial growth in a batch culture over time can be graphed as log (# of cells/ml) (y-axis) versus
time (x-axis). This is called a growth curve. The cell number is plotted as the log of the cell
number, since it is an exponential function. The growth curve has four distinct phases: lag,
exponential (log), stationary, and death phase. The lag phase is characterized by no increase in
cell number, but the bacteria are metabolically active and are adjusting to their new environment.
During an exponential (log) phase, bacteria grow at maximum rate, doubling at a constant rate.
Generation time (time it takes bacteria to double) is also determined at this phase. During
stationary phase, there is no increase in the number of cells, due to exhaustion of nutrients, buildup of wastes, or lack of space. Death phase is the final phase when the media and conditions can
no longer sustain life and the rate of cell growth is less than the rate of cell death. This phase
often can’t be observed using turbidimetry because dead cells can still scatter light. Instead,
CFU counts, which measure viable and culturable cells, can be used to better identify death
phase, as shown in the graph below. Source taken and revised from
www.umsl.edu/microbes/files/pdfs/introductiontobacteria.pdf
There are two ways you can grow a liquid culture. A batch culture refers to a closed system
where microorganisms are incubated with a single batch of medium thus the cells cannot
maintain exponential growth for long. A continuous culture refers to an open system where a
constant circulation of fresh medium and removal of spent medium allows the bacterial
population to maintain a constant doubling time and maintain balanced growth. A common
67
piece of equipment used to grow a continuous culture is called a chemostat.
In today’s lab, you will be working with batch cultures. E. coli will be grown for approximately
12 hours in 6 different conditions:
1. Rich Media LB @ 37°C No Shaking
2. Rich Media LB @ 37°C Shaking
3. Rich Media LB @ 25°C Shaking
4. Minimal Medium @ 37°C Shaking
5. Minimal Medium @ 37°C No Shaking
6. Minimal Medium @ 25°C Shaking
PROCEDURE
1. The first lab class of the day will inoculate 200 ml of media with 4 ml of overnight culture.
2. For the next 12 hours, withdraw 3 ml to measure the OD at 595nm every 30 min.
§ For non-shaking cultures, be careful to swirl the flask to suspend the cells evenly prior to
withdrawing your sample.
§ Note that when the previous OD595 reading was higher than 0.5, you should perform a 1:4
dilution prior to performing the OD595 reading (i.e. add 0.6 ml of culture to 1.8 ml of the
appropriate diluent [LB for LB grown cultures OR water for minimal media grown
cultures] and MIX WELL). The reading must then be multiplied by the dilution factor.
3. Use Excel to plot OD595 vs Time. In your lab notebook, paste your graph of OD595 vs Time
(all 6 conditions on 1 graph, see below left). Also, plot Concentration vs Time (pick any 1
condition and use a standard curve from the Turbidimetry lab, see below right). Lastly,
determine the generation time for each flask by choosing 2 points from exponential (log) phase
and using the equation below.
68
Generation Time Equation:
G= time/number of generations = t/n
n= log (#cells at end) - log (# of cells at start)/ log 2
The above equation simplifies to: n= 3.3 x log (# cells at end/# cells at start)
Post-lab Questions
1. Predict growth curves for the following conditions:
§ LB @ 37 S vs LB @ 25 S [Note: S=shaking; NS=no shaking]
§ LB @ 37 S vs MM @ 37 S
§ LB @ 37 S vs LB @ 37 NS
2. How well do the actual growth curves match the predicted growth curves? Explain.
3. What effect does “no shaking” have on the cells in a batch culture? Is the “no shaking” data
for this experiment truly accurate for a “no shaking” culture? Explain.
69
BACTERIAL MOTILITY
OBJECTIVES: Students should be able to
1. Explain the biological basis of motility and chemotaxis
2. Describe the various arrangements of flagella
3. Set up a hanging drop slide
4. Inoculate motility agar
5. Interpret a hanging drop slide and motility agar reaction
A large number of bacteria are motile. Most possess one or more flagella on their surface that
allow them to swim. The pattern of flagellation is an important feature in identification of motile
bacteria. The figure illustrates some commonly observed arrangements of flagella.
Polar flagella occur at one or both ends or “poles” of the bacterium. There may be a single polar
flagellum (unipolar) or tufts of polar flagella. Amphitrichous flagella are when a single
flagellum is present on both poles of the bacteria (not shown). Lophotrichous flagella are a tuft
of flagella at one or both poles of the bacteria. Peritrichous flagella are distributed around the
surface of the organism. Most motile bacteria move in a straight line for a brief time, then turn
and randomly change directions before swimming again. The straight line movement is called a
run and the turn is called a tumble. Runs and tumbles are controlled by the clockwise or
counterclockwise rotation of the basal body of the flagellum, the motor that is anchored in the
70
cell membrane. Some bacteria do not tumble, but rather reverse direction when they reverse the
rotation of the basal body. Many flagellated bacteria can move toward useful chemicals and
away from harmful ones. This ability to control movement in response to chemical stimuli is
termed chemotaxis. Chemotactic bacteria contain receptors in the cell membrane that bind to
certain chemicals and cause the basal body to direct either a run or tumble (or forward and
reverse directions). When the chemical stimulus is an attractant (or repellant), such as a rich
nutrient source (or a poison), the basal body rotates so that the bacteria swim in straight lines
toward (or away from) the signal for long periods of time. If no stimulus is present, the basal
body reverses direction, causing the bacterium to tumble more often.
Flagella can be visualized by a somewhat difficult staining technique, but their presence is
typically based on detecting motility of living bacteria. Observations can be made in a hanging
drop and using semisolid motility agar.
HANGING DROP SLIDE
This method is commonly used to view living organisms for the rapid determination of motility.
The hanging drop is prepared by suspending a fluid sample from a coverslip over a depression
well in a specially designed microscope slide. Wet mounts can be used for the same purpose,
however, wet mounts tend to dehydrate rapidly. Hanging drops, on the other hand, are sealed
within the depression and retain their liquid for longer periods of time. In both methods, the
living specimen is unstained. For best results, reduce the amount of light passing through the
specimen.
MATERIALS
•
•
•
•
•
Depression slides
Coverslips (glass)
Petroleum jelly
Toothpicks
Cultures (in broth):
• Positive Control: Proteus vulgaris or Escherichia coli
• Negative Control: Staphylococcus epidermidis
PROCEDURE
1. With the toothpick, add a small amount of petroleum jelly to the depression slide where
the edges of the coverslip will sit (see diagram below). This will help the coverslip adhere
to the slide.
2. Place one or two loopfuls of a culture in the center of the coverslip.
3. Place the coverslip on the depression slide with the depression under the drop of fluid.
4. Examine with low power. Focus on the edge of the drop.
71
5. Switch to high power. These slides are thicker than most microscope slides so be careful.
You may add some immersion oil to the coverslip and change objective lenses. Some fine
adjustment may be necessary, but do not turn the coarse adjustment. Look for motility
and be sure to distinguish it from Brownian motion. True motility results from flagellar
rotation.
6. Make sure to return your depression slide to the TA after use.
MOTILITY AGAR
Motile bacteria require liquid to move. Thus bacteria can propel themselves in broth or across
the surface of a wet agar plate. They will not however move when embedded in 1.5% agar, the
minimum concentration found in most agar media. Semisolid agar has a reduced agar
concentration (0.4%) that allows flagellated bacteria to migrate from the site of inoculation.
Semisolid media are prepared in tubes and are inoculated through most of their length by
stabbing with a needle. Thus after 48 hours of incubation, growth of a motile organism will be
observed as a turbid region extending from the stab. Nonmotile bacteria will only grow along
the stab line.
MATERIALS
•
•
Semisolid nutrient agar in tubes
Cultures (in broth):
• Positive Control: Proteus vulgaris or Escherichia coli
• Negative Control: Staphylococcus epidermidis
PROCEDURE
1. Using aseptic techniques, stab the needle into a broth culture, then inoculate the motility
agar tube by stabbing with the needle to approximately three-quarters of its depth. Be
careful to stab into the center of the medium and not to touch the side of the tube.
2. Incubate at room temperature for 48 hours or at 37° C overnight.
3. Examine tubes after incubation.
72
Post-lab Questions
1. What advantages might motile bacteria have over nonmotile bacteria?
2. How could you experimentally demonstrate positive chemotaxis?
73
First group project:
INVESTIGATION OF AN ENVIRONMENTAL SAMPLE
One of the most important skills in science is the ability to design a logical, well-crafted
experimental study. During the first group project, you will gain practice in developing an
experimental plan while applying methods you learned in the first part of the course.
à Your goal in this project is to “sample” an environment and identify the bacteria in your
sample, using the techniques you have learned thus far. You will work in groups to accomplish
the project and present your results to the class in a scientific poster.
Preparation
Choose an environment with a variety of microbes for sampling. Your TA must approve your
selection. Some examples of good environments include:
• Soil, sand, or sediment
• Ocean, pond, or lake water
• Organismal contents of invertebrates such as worms or insects
• Resources from vertebrates such as cats and dogs
• Plants or seaweed
First group project plan (See syllabus for due date)
1. The experimental plan should include the following:
• The environment you are sampling, and
• A hypothesis stating four different species of bacteria that you expect to find in your
sample. At most, only one of the species chosen can be bacteria found in the lab manual.
The four species you predict need to be supported with at least four primary
literature articles.
• Include screenshots of the first page of those 4 primary literature articles.
2. List the methods that you will use to identify the bacteria in your sample.
• Required methods are to do a gram stain and observe morphology.
• Three additional methods must be included in the plan. Methods you have learned so far
in this course include microscopy and staining techniques, media plates, anaerobic
culture techniques, and biochemical tests.
3. Carefully and thoughtfully (minimizing waste), request the materials you need to carry out
your project.
Preparing the first group project poster (See rubric posted on Canvas)
1. Generate a dichotomous key for your project. The hypothesized and actual dichotomous key
should be included in the poster. They could be color coded into one figure.
2. Draft the introduction and methods sections of your poster. From the rubric:
Introduction
o Background on where/how sample was acquired
o Background on why the chosen environment allows growth of different bacteria
o Background on possible types of bacteria present
74
o Primary literature cited
Methods
o All methods used are listed
o How methods were performed is explained; standard procedures from the lab manual
(i.e. Gram stain) should not be described
3. Draft the Results and Discussion sections of your poster. From the rubric:
Results
o Tables and graphs are in the correct format
o Tables, graphs, and additional results not in “picture format” are stated
Discussion
o Significance of results is discussed – what each test tells you about the properties of
the bacteria
o Possible genera (and species if you have enough information to make an assessment)
are stated
o The connection between the results and the possible genera
o Whether your hypotheses are supported or not is explained
o Possible sources of error
4. Format your poster
o Poster templates are available on Canvas
5. Practice presenting the poster as a group
75
Biol302(Group(Project(Plan(Schedule
Group&name:&__________________________________________
Section&Number:&_______________________________________
Request&date:
Experiment
Materials
Experiment
Materials
Experiment
Materials
1
Start&date:
2
3
4
5
Request&date:
1
Start&date:
2
3
4
5
Request&date:
1
Start&date:
2
3
4
5
This document may be helpful for planning but does not need to be turned in. See the previous
page for the project plan guidelines.
76
Poster Presentation Rubric
Group_____________________
Poster
Abstract
Provides a clear summary of the entire study
Hypothesis
Stated where necessary & complete
Introduction
Background on where/how sample was acquired
Background on why the chosen environment allows growth of different bacteria
Background on possible types of bacteria present
Primary literature cited
Methods
Dichotomous key included
All methods used are listed
How methods were performed is explained; standard procedures from the lab manual (i.e. Gram
stain) should not be described
Results
Tables and graphs are in the correct format
Tables, graphs, and additional results not in “picture format” are stated
Dichotomous key results included or results indicated in methods dichotomous key.
Discussion
Significance of results is discussed – what each test tells you about the properties of the bacteria
Possible genera (and species if you have enough information to make an assessment) are stated
The connection between the results and possible genera (refer to dichotomous key)
Whether your hypotheses are supported or not is explained
Possible sources of error
References
Minimum 4 primary literature articles
Articles listed in the correct format
Required elements are included
Gram+/-, morphologies, and 3 other methods
Poster formatting
Information flows from one section to the other
Poster is aesthetically pleasing
If printed out, font large enough to be read from approx. 4 feet away
5-10 minute presentation
Delivery
Volume of talk
Speed of talk
Stayed within time limit
Coverage
Each section of the poster was discussed (except for abstract, which should not be covered during
presentation)
All results were stated
Significance of results
Participation
All group members participated equally
Questions
Questions asked were answered correctly and completely
Group members participated in answering questions equally
77
QUORUM SENSING
OBJECTIVES: Students should be able to
1. Describe how quorum sensing controls gene expression in bacteria.
2. Explain the role of autoinducer in quorum sensing.
3. Explain how and when Vibrio harveyi can produce light.
Since bacteria are unicellular organisms, for many years it was assumed that each bacterial cell
functioned independently of other cells, and that cell-cell communication was common only
within multicellular organisms. The realization that bacterial cells can communicate with each
other and exhibit behaviors as a community became widely acknowledged when the molecular
mechanisms behind quorum sensing began to be discovered in the 1990’s. Quorum sensing
refers to the idea that a community of bacteria senses the number of other bacteria present to
control gene expression in the members of the bacterial community.
Quorum sensing was first described by scientists studying bioluminescence in marine bacteria
(Vibrio fischeri) that forms a symbiosis with certain species of squid and fish. Since the
discovery of quorum sensing in V. fischeri, quorum sensing appears to be performed by virtually
all species of bacteria, controlling important processes such as pathogenesis, gene transfer, and
biofilm formation.
The quorum sensing system of V. fischeri is unusually simple. The lux operon, which encodes
the light-producing enzyme luciferase, is transcriptionally activated by the LuxR protein when
bound to a small molecule called the autoinducer. For gram-negative bacteria, the autoinducer
is usually a type of homoserine lactone. In V. fischeri, synthesis of the autoinducer requires the
LuxI enzyme. The bacteria constitutively synthesize autoinducer, which freely diffuses into and
out of bacterial cells. When the bacterial population is at a high density (such as the density
reached within the confined space of a squid’s light organ), the concentration of autoinducer is
high enough so that many molecules of autoinducer-LuxR complex will bind to DNA and
activate transcription of the lux operon. Thus, V. fischeri produces light only when the
population density is high.
For most other bacteria, such as Vibrio harveyi, quorum sensing is not controlled by a simple
system involving LuxR. Instead, quorum sensing is controlled by two-component signaling
pathways that sense the autoinducer and transduce the signal to control transcription. V. harveyi
actually senses two different autoinducer molecules using two different signaling systems. The
first system senses autoinducer (AI-1) produced by V. harveyi whereas the second system senses
autoinducer (AI-2) produced by other species of Gram-negative bacteria. Thus, V. harveyi can
participate in both intraspecies and interspecies communication.
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In this lab, you will add different volumes of “cell-free supernatant” (CFS) to fresh cultures of V.
harveyi and observe bioluminescence. The CFS is prepared by culturing V. harveyi in liquid
medium overnight, then separating the cells from the supernatant by centrifugation.
MATERIALS
•
•
•
•
overnight culture of V. harveyi in AB medium
fresh AB medium
1 microcentrifuge tube
3 glass culture tubes
PROCEDURE
Visual overview of procedure:
This experiment will be performed with your lab group. Label the microcentrifuge tube and 3
glass cultures tubes with your group number. Also, label the 3 GLASS culture tubes as follows:
• Media only
• Cells+Media
• Cells+CFS
1. Prepare the cell free supernatant (CFS) using the microfuge tube.
• Using aseptic technique, pipet 700 ul of the overnight culture into the microcentrifuge
tube and centrifuge at maximum speed (>13,000 x g) for 5 min to pellet the cells.
• Transfer 500 ul of the supernatant to a microfuge tube labeled “CFS.” Be careful to
avoid the cell pellet, which you will need for the next step.
2. Use the pellet left over from step 1 to make a cell suspension.
• Remove the remaining supernatant from the microfuge tube using a micropipette and
discard as culture waste.
• Add 500 ul of fresh AB medium to the cell pellet and pipet up and down to resuspend
until no clumps are visible. This is your “cell suspension”.
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3. Set up your new cultures. Use aseptic technique.
• Pipet fresh AB medium into each GLASS tube:
o 1 mL in the “Media only” tube
o 800 ul in the “Cells+Media” tube, and
o 500 ul in the “Cells+CFS” tube.
• Then, add 200 ul of the cell suspension to each of the following tubes:
o “Cells+fresh media”
o “Cells+CFS”
• Finally, add 300 ul CFS from the CFS microfuge tube to the “Cells+CFS” glass tube.
4. Place the 3 glass tubes “Media only,” “Cells+fresh media,” and “Cells+CFS” on the shaker.
Note the time.
5. After 1 hour and 2 hours on the shaker, observe your tubes next to the “Media only” tube in a
dark room. It may take up to 5 minutes for your eyes to fully adjust to the dark; try not to agitate
your tubes in the meantime. Once your eyes have adjusted, swirl the tubes in the darkroom and
observe the tubes for a minute after you have stopped swirling.
6. Record your observations in your notebook. At the end of the lab, place your tubes in the tube
waste.
Post-lab Questions
1. What were the expected results? Explain the rationale of the experiment. If you didn’t obtain
the expected results, hypothesize why not and describe how the experiment could be improved.
2. What are differences between luminescence and fluorescence?
3. Why do V. harveyi cultures glow more brightly when swirled vigorously? Be specific.
4. Imagine you have a mutant V. harveyi strain that can’t synthesize autoinducer and another
mutant V. harveyi strain that makes a defective luciferase. If you streaked both strains near (but
not touching) each other on the same plate, would you expect to see light produced? If so, by
which strain, where, and why?
5. You have two identical tubes of wild-type V. harveyi (with the same cell concentration in
each tube) in fresh media. To one tube, you add CFS from a light-producing V. fischeri culture,
and to the other tube you add an equal volume of fresh media. If both tubes of V. harveyi have
the same light production, what can you conclude? Explain.
6. Many biomedical research labs study quorum sensing in V. fischeri or V. harveyi. How is
studying light production in marine bacteria relevant to human health? List 3 different examples.
7. Why do you think it is important for bacteria to be able to control the expression of some
genes based on the number of individuals present?
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Dichotomous Key for Bacterial Identification
Based on the characteristics/properties of the organisms, create a key to successfully
differentiate and identify each of these organisms from a mixed culture containing all of
them.
Neisseria meningitidis
Corynebacterium xerosis
Staphylococcus aureus
Lactobacillus delbrueckii
Staphylococcus epidermidis
Proteus vulgaris
Bacillus megaterium
Escherichia coli.
Use your lab manual, experiments you have conducted in class, your text book, and
Bergey’s manual as resources to help you design your key.
Example: How to set up your key
Mixed Culture
(Bacteria A, B, C, and D)
Test 1
Positive Reaction
A, C, and D
Negative Reaction
B
Test 2
Positive Reaction
D
Negative Reaction
A and C
Test 3
Positive Reaction
C
Negative Reaction
A
Remember, in place of a “test” you can use a property like morphology, gram stain,
capsule stain, motility, biochemical tests, selective media, etc. “Positive” and “Negative”
reaction can be rod or cocci (if using morphology), capsule present or absent, etc.
Include a list of the sources you used. You do not need a formal citation, just name the
text or website and the page you used or the date you went to the website.
Important Notes:
• This assignment must be completed individually. Since there are so many possible tests
available, each student’s dichotomous key should be substantially different from that of other
students.
• Include a list of the sources you used. You do not need a formal citation, just name the text or
website and the page you used or the date you went to the website. Include a reference for each
test.
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BIOCHEMICAL REACTIONS
OBJECTIVES: Students should be able to
1. Explain the biological basis of each biochemical reaction
2. Perform each biochemical test and interpret the test reactions
3. Apply these tests to the identification of an unknown
4. Describe some common multiple test and rapid test systems used in bacterial
identification
5. Compare the advantages and disadvantages of immunological assays and DNA probes
6. Explain the polymerase chain reaction and its use in microbiology
Bacteria are among the most diverse organisms with respect to the types of biochemical and
metabolic reactions they can perform. You will be using some common biochemical tests in the
identification of your unknown. A variety of specialized media is used to investigate the
biochemical reactions of bacteria. Make use of the DIFCO manual for the composition of the
media.
FERMENTATION REACTIONS
Sugars
Most bacteria obtain energy from the oxidation of organic compounds such as sugars. Strict
aerobes oxidize sugars to carbon dioxide and water through the process of aerobic respiration,
using a cytochrome electron transport system and molecular oxygen as the final electron
acceptor. Many strict anaerobes perform fermentation reactions. They oxidize and then reduce
sugars to organic compounds (acids, alcohols, aldehydes). No cytochrome system is used in
fermentations and oxygen is not involved. Fermentation is not as efficient an energy-yielding
process as aerobic respiration. Facultative anaerobes possess the enzymes that allow them to
perform either respiration or fermentation depending upon the availability of oxygen.
The types of compounds that an organism can ferment and the end products that are produced are
genetically determined. Among the end products commonly generated by fermentation are
organic acids and gases, such as carbon dioxide and hydrogen.
Fermentation test media contain:
1. a single carbohydrate as an energy source
2. nonfermentable sources of nitrogen and other nutrients
3. a pH indicator such as phenol red or bromocresol purple
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4. a Durham tube (an inverted vial) that collects gas
A positive fermentation reaction is indicated by a change in the color of the indicator from red
(or purple) to yellow, below pH 7, due to acidic fermentation products. Gas production, if
observed, is also recorded. Uninoculated controls must be performed in order to accurately
evaluate results.
Examples of fermentation tubes after incubation (left to right): red, no gas; yellow, gas; yellow,
no gas
Expected results with species in this lab:
Streptococcus faecalis
Glucose
+
Sucrose
+
Mannitol
+
Pseudomonas putida
+
-
Escherichia coli
+ and gas
+ and gas
MATERIALS
•
•
Fermentation broths:
• Glucose
• Sucrose
• Mannitol
Cultures (broth):
• Escherichia coli
• Pseudomonas putida
• Streptococcus sp.
PROCEDURE
1. Inoculate each broth with a loopful of the organism. Ensure that the caps are screwed on
tightly. Incubate for 24 - 48 hours at 37°C. A set of negative control tubes
(uninoculated) must be kept for comparison.
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2. Examine the tubes for growth (+), acid (A) and gas production (G) and compare to the
control tubes. Record your results.
Post-lab questions
1. If organisms do not grow in fermentation media, what should your interpretation be?
2. Explain why some organisms grow in the media but do not produce acid.
Methyl Red and Voges-Proskauer Tests (MR-VP)
These fermentation tests are used to differentiate between certain intestinal bacteria called
coliforms. The MR-VP broth contains dextrose as the carbohydrate source. Some coliforms
will ferment the dextrose to acid products that will cause the pH to drop below pH 5. This is
called a mixed acid fermentation. After incubation, the addition of methyl red dye, which
turns red below pH 4.4, will indicate whether such fermentation has occurred. Other coliforms
will convert dextrose to products such as ethanol or butanediol, which are not acidic. These
bacteria are negative in the methyl red test and the media remains yellow.
Butanediol fermentation is demonstrated by the Voges-Proskauer test, which measures the
presence of acetoin (acetyl methyl carbinol), a precursor to butanediol. This test uses the same
medium as the methyl red test and both tests are usually performed in parallel. Barritt's reagents
(alpha-naphthol and potassium hydroxide) are added to a 48 hour culture, and the tube is shaken
to aerate the solution. The development of a pink or red color after agitation is a positive
reaction for the production of acetoin.
MATERIALS
•
•
•
MR-VP broth
Methyl red and Barritt’s reagents (A & B)
Cultures
• Escherichia coli (positive for MR; negative for VP)
• Enterobacter aerogenes (positive for VP; negative for MR)
PROCEDURE
1. Inoculate the organism. Incubate for 48 hrs or 5 days at 37°C. (A single uninoculated
control should be kept.)
2. Remove 1 ml from each culture to a clean tube for the VP test.
3. Methyl red test: Add a few drops of methyl red to the original culture tube and mix the
contents. Record your results.
4. Voges-Proskauer test: Add 0.5 ml (~15 drops) of Barritt's reagent A to the tube and mix.
84
Add 0.5 ml of Barritt's reagent B and mix. Aerate the tube by mixing occasionally over a
one hour period or until a pink or red color develops. Record your results.
ENZYME PRODUCTION
Among the many enzymes that bacteria may produce are exoenzymes, enzymes that are
excreted, and are used to degrade large polymers into smaller compounds. The detection of such
enzyme activities is often confirmatory in identification of unknowns. For example, starch
digestion results from the action of amylase released into the surrounding medium. The starch is
a polysaccharide that cannot pass across the cell membrane. Amylase breaks starch into smaller
sugar residues that can enter the cell and be processed by respiration or fermentation. Gelatinase
is another exoenzyme. It can cause the liquefaction of media solidified by gelatin (rather than
agar). Caseinase is an enzyme that hydrolyzes casein, the major protein component in milk. As
a result of proteolysis, breakdown of protein, by the enzyme, milk incorporated into agar
medium loses its characteristic white appearance and becomes transparent. Lipase production is
common to bacteria that grow in foods rich in fats such as butter and mayonnaise. This enzyme
breaks fats into its components glycerol and fatty acids. Agar media for detecting lipase activity
contains lipids prepared from egg yolks. The media loses its opacity in the area surrounding a
lipase-producing bacterium.
Most enzymes are endoenzymes. They are produced in the cell and catalyze intracellular
reactions. Examples of reactions, used to identify unknown bacteria, that are catalyzed by
endoenzymes are a) the breakdown of toxic wastes such as hydrogen peroxide or urea, b) the
reduction of nitrate or oxygen, c) the degradation of specific amino acids, and d) the utilization
of noncarbohydrate carbon sources for growth.
Amylase Production
Amylase activity is demonstrated using starch agar, a medium containing starch as the
carbohydrate source. After growing bacteria on the starch agar, the plate is covered with Lugol’s
Iodine, which binds to starch. If Amylase is present, starch in the media near the bacteria will be
degraded, and there will be a clear zone in the media. If Amylase is absent, starch is not
degraded and there will be no clear zone in the media.
MATERIALS
•
•
•
Starch agar plates
Lugol’s Iodine
Cultures
• Positive Control: Bacillus subtilis
• Negative Control: Escherichia coli
PROCEDURES
1. Streak each organism across a small portion of the starch agar surface.
85
2. Incubate at 37°C for 48 hours.
3. Cover the surface with Lugol’s iodine. Rotate to distribute the iodine into a thin layer. Do
not flood the plate.
4. Record your results.
Catalase production
Catalase is an enzyme that detoxifies hydrogen peroxide, a compound that is a byproduct of
aerobic respiration and is lethal if it accumulates in the cell. Catalase breaks down the hydrogen
peroxide into water and oxygen. All respiring organisms therefore must have some mechanism
for detoxification, and catalase is one of the common methods. When hydrogen peroxide is
added to a colony of catalase-producing bacteria, it is broken down into water and oxygen, and
the oxygen produced can be seen as bubbles. Remember that not all organisms that live in
oxygen have catalase; aerotolerant anaerobes are indifferent to the presence of oxygen, since
they ferment in the presence or absence of oxygen.
MATERIALS
•
•
•
TSA plates
3% hydrogen peroxide
Cultures (agar slant):
• Positive Control: Klebsiella pneumoniae
• Negative Control: Streptococcus sp.
PROCEDURES
1. Streak each organism across a small portion of the TSA surface.
2. Incubate at 37°C for 48 hours.
3. Place a few drops of 3% hydrogen peroxide over a colony.
4. Observe whether oxygen is produced.
86
Post-lab question
1. What happens to obligate anaerobic organisms in the presence of oxygen? Why?
Oxidase Production
Oxidases are enzymes that catalyze the reduction of oxygen during respiration. For example, in
most Gram-positive bacteria and many Gram-negative bacteria, Cytochrome Oxidase performs
the final step in the electron transport chain, reducing oxygen to water. Other bacteria, such as
those in the Enterobacteriaceae family, do not reduce oxygen using this enzyme. Thus, detection
of Cytochrome Oxidase is a valuable tool in differentiating among bacteria. The test utilizes a
colorless reagent to detect Cytochrome Oxidase. In the presence of oxygen and Cytochrome
Oxidase enzyme, the oxidase reagent forms a pink/maroon/dark blue-black compound.
MATERIALS
•
•
•
Sterile cotton swab
Oxidase reagent
Cultures (agar slant):
• Positive Control: Pseudomonas putida
• Negative Control: Escherichia coli
PROCEDURES
1. Obtain a small sample of microbe on the tip of a sterile cotton swab.
2. Place two drops of oxidase reagent over the sample.
3. Observe the color change. A positive reaction may appear pink at first, then maroon and
finally black/dark blue. Take care to avoid contact with the oxidase reagent. NOTE: An
alternate procedure is performed by placing some oxidase reagent directly on the colony
on the agar or on organisms applied to a filter disk.
Post-lab question
87
1. How is ATP generated as a result of electron transport systems?
Tryptophan hydrolysis (Indole Production)
The ability to degrade amino acids to identifiable end products is often used to differentiate
among bacteria. Tryptophan, for example, is hydrolyzed to indole, pyruvic acid and ammonia by
tryptophanase. The pyruvic acid can be further metabolized to produce large amounts of energy.
The ammonia is available for use in synthesis of new amino acids.
Indole can be detected with Kovac's reagent (para-dimethylaminobenzaldehyde in isoamyl
alcohol). The para-dimethylaminobenzaldehyde reacts with the indole in the media to form a red
dye, which forms a complex with the isoamyl alcohol, producing a red layer at the top of the
tube.
MATERIALS
•
•
•
1% Tryptone broth
Kovac’s reagent
Cultures (broth):
• Escherichia coli (positive control)
• Enterobacter aerogenes (negative control)
PROCEDURES
1. Inoculate tryptone broth (1%) with the organism.
2. Incubate at 37°C for 48 hours.
3. Add 10 drops of Kovac's reagent. A red color in the alcohol (upper) layer is a positive
result.
Post-lab question
Why do you think it is important not to incubate your cultures for more than 5 days when
performing the indole test?
88
Citrate Utilization
Some bacteria may be able to use organic compounds other than sugars as their sole source of
carbon. For example, the ability to metabolize citrate is useful for differentiating among
Enterobacteriaceae. Simmons Citrate agar is a medium containing citrate as the sole carbon
source and ammonium salts as the sole nitrogen source. Citrate utilization produces an alkaline
carbonate, increasing the pH of the medium. Bromothymol blue is present in the medium as the
indicator dye. It is green at neutral pH (if citrate was not utilized) and deep blue above pH 7.6 (if
citrate was utilized).
Koser's citrate broth is another medium used to test for citrate utilization. Growth is evidence of
a positive reaction.
MATERIALS
•
•
Simmons citrate agar
Cultures (broth):
• Enterobacter aerogenes (positive control)
• Escherichia coli (negative control)
PROCEDURES
1. Using a sterile inoculating needle, streak one organism over the surface of the agar slant,
then stab the butt. Repeat with the second organism.
2. Incubate the tubes at 37°C for 48 hours.
3. Examine the tubes. Is there a color change in the media?
Post-lab question
Why does the media become alkaline when bacteria grow on Simmons citrate agar?
89
Single Media/Multiple Tests
Several media are designed to yield more than one biochemical reaction. Among the more
commonly used media in this category are SIM media, Triple Sugar Iron agar (TSI) and Kliger's
Iron agar (KIA). SIM medium derives its name from three reactions: production of hydrogen
sulfide from sulfur-containing amino acids, indole production and motility. Motility is observed
if bacteria migrate from the stab line throughout the semisolid medium. Blackening of the
medium indicates production of hydrogen sulfide, since the hydrogen sulfide reacts with ferrous
ammonium sulfate and forms ferrous sulfide (a black precipitate). After the addition of Kovac’s
reagent, a red color indicates the presence of indole. The medium is used primarily for
differentiation of gram negative enteric bacteria.
MATERIALS
•
•
SIM agar
Cultures (broth):
• Escherichia coli
• Pseudomonas putida
• Salmonella typhimurium
PROCEDURES
1. Inoculate the SIM agar using a needle by a single stab through the center to about ¾ the
length of the agar.
2. Incubate E. coli and S. typhimurium at 37°C for 48 hours. Incubate P. putida at RT or
30C.
3. Examine the tubes for growth and color change.
4. Add a few drops of Kovac’s reagent to the SIM tubes and observe for a red color change,
indicating indole production.
Triple sugar iron agar (TSI) or Kliger's iron agar (KIA) and are widely used in the
identification of gram negative bacteria particularly the Enterobacteriaceae. The media are
identical except that TSI contains sucrose in addition to the dextrose (also known as D-glucose)
and lactose found in KIA. The media are poured as slants and are inoculated with a stab to the
butt followed by a streak of the slant surface. The bacteria therefore are exposed to both an
anaerobic environment (butt) and an aerobic one (slant). Phenol red is present as an indicator.
Do not tighten the cap on the tube.
If the bacteria are nonfermenters, such as Pseudomonas, they can grow on the slant by the
aerobic degradation of peptides in the medium to alkaline products. The slant and the butt will
remain red.
If the bacteria can ferment dextrose, but not sucrose or lactose, acid is produced in the slant and
90
the butt and the medium turns yellow. However, there is only a low concentration of dextrose
(also known as D-glucose) in TSI media (0.1% glucose compared to 1% lactose and 1%
sucrose), so the dextrose is used up within 12 hours. Bacteria at the surface continue to grow by
degrading peptides. By 18 to 24 hours, the alkaline end products cause the medium in the slant
to revert to a red color. Such reactions are characteristic of Shigella and other nonlactose
fermenters.
If the bacteria can ferment lactose and/or sucrose as well as dextrose (also known as Dglucose), the slant and butt will remain yellow after prolonged incubation. The high
concentration of lactose and/or sucrose breakdown products keeps the slant acidic despite the
production of alkaline products by protein degradation.
TSI and KIA also contain sodium thiosulfate and ferrous sulfate as indicators of hydrogen
sulfide production. Salmonella sp. (dextrose fermenters, lactose nonfermenters) will yield an
acid butt with a black precipitate and an alkaline slant.
MATERIALS
•
•
TSI slants
Cultures (broth):
• Escherichia coli
• Salmonella typhimurium
• Pseudomonas aeruginosa
• Shigella boydii
PROCEDURES
91
1. Inoculate the TSI using a needle by a single stab through the butt of the agar and
streaking the surface of the slant.
2. Incubate the cultures at 37°C (E. coli, S. typhimurium and S. boydii) or 30°C (P. putida)
for 48 hours.
3. Examine the tubes for growth and color change.
92
SUMMARY OF BIOCHEMICAL REACTIONS
FERMENTATION REACTIONS:
1. Sugars:
-Sugar oxidizes to organic compounds: acids, alcohol, or aldehyde (many strict
anaerobes)
-Sugar oxidizes to: H2 or CO2 (strict aerobes)
-If test is positive, red/purple =>yellow (below pH 7)
-If gas was produced, Durham tube will collect the gas
Organisms: E. coli, P. putida, Streptococcus sp.
1. Inoculate each fermentation broth (glucose, sucrose, mannitol) with a loopful of the
organism.
2. Incubate for 24-48 hours at 37°C. Record observations next lab period.
2. Methyl Red (MR-VP):
-Dextrose (D-Glucose) sugar oxidizes to acid products or neutral alcohol products.
-If test is positive, yellow =>red (below pH 4.4) after addition of methyl red
Control organisms: E. coli (positive for MR; negative for VP); E. aerogenes (positive for VP;
negative for MR)
1. Remove 1 ml from each culture (that was inoculated last time) to a clean tube and save it
for the VP test.
2. Add a few drops of methyl red to the original culture tube and mix the contents.
3. Record your results on the same day.
3. Voges-Proskauer Tests (MR-VP):
-Dextrose (D-Glucose) sugar oxidizes to acid products or intermediate compound
called acetoin during the production of the neutral end product, butanediol.
-If test is positive then should turn yellow =>red (presence of acetoin) after an
addition of Barritt’s reagents.
Control organisms: E. coli (positive for MR; negative for VP); E. aerogenes (positive for VP;
negative for MR)
1. To the saved 1 ml of each culture, add 15 drops of Barritt’s reagent A then 15 drops of
Barritt’s reagent B to the original culture tube and mix the contents.
2. Record your results on the same day.
93
ENZYME PRODUCTION:
1. Amylase Production:
-Amylase breaks starch into smaller single subunits of alpha D-glucose
-If test is positive, there will be a clear zone on the agar surface (due to digestion)
after addition of Lugol’s iodine.
-If test is negative, there will be a purple/blue zone on the agar surface after addition
of Lugol’s iodine.
Positive control organism: Bacillus subtilis/Bacillus megaterium
Negative control organism: E.coli
1. Streak each organism across a small portion of the starch agar surface.
2. Incubate at 37°C for 48 hours.
3. During the next lab period, cover the surface with Lugol’s iodine. Rotate to distribute
the iodine into a thin layer. Do not flood the plate. Record your result.
2. Catalase production:
-Catalase splits toxic 2H2O2 => 2H2O + O2
- If test is positive, there will be oxygen bubbles after addition of H2O2
Positive control organism: Klebsiella pneumoniae
Negative control organism: Streptococcus sp.
1. Streak each organism on one half of the TSA plate.
2. Incubate at 37°C for 48 hours.
3. During the next lab period, cover the surface with few drops of 3% hydrogen
peroxide. Is oxygen produced? Record observations.
3. Oxidase production:
-Cytochrome oxidase is the terminal protein in the Electron Transport Chain and is
involved in reducing O2 to H2O
-Enterobacteriaceae do not use this enzyme to reduce O2 to H2O
- If test is positive, sample changes from colorless to pink => maroon => black after
an addition of oxidase reagent
- If test is negative, sample remains colorless after an addition of oxidase reagent
Positive control: P. aeruginosa (on agar)
Negative control: E. coli
1. Obtain a small sample of microbe on the tip of a sterile cotton swab.
2. Place two drops of oxidase reagent over the sample.
3. Observe the color change and record.
94
OTHER BIOCHEMICAL REACTIONS:
1. Citrate utilization:
- Some bacteria can metabolize citrate as their carbon source instead of sugars.
- Bacteria that metabolize citrate use ammonium salts and release ammonia thus
increasing the pH of the medium
- Useful for differentiating Enterobacteriaceae
- If test is positive, there will be growth and blue color on Simmon’s citrate slant
agar.
- If test is negative, no growth and no color change on Simmon’s citrate slant agar.
Positive control: Enterobacter aerogenes
Negative control: E. coli
2 tubes per group
1. Using a sterile inoculating needle, streak one organism over the surface of the agar
slant, then stab through the butt. Repeat with the second organism.
2. 2.Incubate the tubes at 37°C for 48 hours.
2. Indole production (Tryptophan hydrolysis):
-Tryptophanase enzyme can hydrolyze/degrade tryptophan to indole, pyruvic acid,
and ammonia.
-If test is positive, should see dark red color in the alcohol (upper) layer after an
addition of Kovac’s reagent
-If test is negative, faint pink but no dark red color in the alcohol (upper) layer after
an addition of Kovac’s reagent
Positive control: E. coli
Negative control: Enterobacter aerogenes
2 tubes per group
1. Inoculate tryptone broth (1%) with the organism. Incubate at 37°C for 48 hours.
2. During next lab period, add 10 drops of Kovac's reagent.
95
SINGLE MEDIA/MULTIPLE TESTS:
1. Sulfide-Indole-Motility (SIM):
-For differentiation of gram negative enteric bacteria
-Three tests:
A. Sufide production: indicated by a blackening of the medium
-If test is positive, black ppt due to lead nitrate already in the agar
-If test is negative, no change in color
B. Indole production is determined after addition of Kovac’s reagent
-If test is positive, should see dark red ring in the alcohol (upper) layer
after an addition of Kovac’s reagent
-If test is negative, faint pink or no color in the alcohol (upper) layer
after an addition of Kovac’s reagent
Organisms: E.coli, P. putida, and S. typhimurium
1. Inoculate anaerobic part (butt) of SIM with the organism.
2. Incubate at 37°C for 48 hours.
3. During the next lab period, add 10 drops of Kovac's reagent and observe for indole
production.
C. Check for motility by observing migration of bacteria in semisolid
medium
-If test is positive, growth throughout medium
-If test is negative, growth around the original stab
2. Triple Sugar Iron agar (TSI):
-Used to identify Gram (-) bacteria/Enterobacteriaceae
- Phenol Red as indicator for sugar fermentation
A. Nonfermenters: slant and butt=red due to utilization of the peptides in the
media.
B. Dextrose (D-Glucose) fermention but NOT sucrose or lactose
fermentation: initially slant and butt=yellow. But by 18-24 hours, slant
becomes red due to utilization of the peptides in the media.
C. Dextrose (D-Glucose) fermentation AND lactose and/or sucrose
fermentation: slant and butt=yellow even after prolonged incubation. Due to
the high concentration of lactose and sucrose, utilization of these sugars
produce acidic products that keep the pH of the media low even if peptides in
the media are also used.
-Sodium thiosulfate and ferrous sulfate as indicators of hydrogen sulfide production
-Black precipitate=H2S production
96
BIOFILM FORMATION AND QUANTITATION
OBJECTIVES: Students should be able to
1. Describe how biofilms are formed and compare the advantages and disadvantages
of bacterial life in a biofilm to that of planktonic bacteria.
2. Grow a biofilm and quantify the biofilm-forming ability of different strains.
3. Develop a hypothesis to explain your observations about biofilms.
Most of the exercises and experiments you will perform during this semester use pure cultures of
free-living bacteria growing in liquid or solid media. In nature, however, bacteria often grow as
populations attached to surfaces in complex structures called biofilms. Biofilms are aggregates
of bacteria encased in a structured exopolymeric matrix that attaches the community to a surface.
Biofilms may be pure cultures consisting of a single type of organism or, more commonly in
nature, a mixed community of organisms. Almost any surface is susceptible to biofilm
formation. Some examples include rocks in a stream, teeth in the mouth, a catheter in the arm, or
pipes delivering water. There are several advantages and challenges to microorganisms that
develop and exist within a biofilm.
Biofilm formation begins when free swimming, planktonic bacteria encounter a surface. The
bacteria become loosely attached to the surface, then form microcolonies. A polymeric matrix
(extracellular polysaccharides, proteins, and DNA) is secreted, encasing the bacteria. These
trapped sessile bacteria form a community that controls the structural complexity of the biofilm.
Some bacteria may escape the biofilm, depending on environmental conditions. Furthermore,
the bacteria may degrade the polymeric matrix using lyase enzymes, to release nutrients the
bacteria can use during harsh times.
The biofilm is not simply a thick layer of extracellular material. There are channels and
pathways that provide access to the interior portions of the biofilm. For more information about
biofilms you may access Biofilms: City of Microbes (review article) and Center for Biofilm
Engineering. Advantages of living in a biofilm include protection from drying (dessication), UV
radiation, antibiotics and other chemicals, and removal from the surface. The channels in the
biofilm can help to trap nutrients. The close cell-to-cell proximity when living in a biofilm
allows cell-cell communication and increased frequency of lateral gene transfer, especially via
conjugation.
In this exercise, you will have the opportunity to grow a biofilm and measure the biofilmforming ability of two different strains of Sinorhizobium meliloti, a soil-dwelling bacterium that
forms a nitrogen-fixing symbiosis with legume plants such as alfalfa.
97
MATERIALS
•
•
•
•
•
•
•
•
6-well plate (1 per lab group)
Rhizobium Defined Medium (RDM)
Overnight broth cultures of S. meliloti in RDM
o Wild-type S. meliloti
o a mutant strain of S. meliloti
Ziploc sandwich bag
LB plate (1 per lab group)
Filtered 0.1% Crystal Violet
distilled water
95% Ethanol
PROCEDURES
Ensure that you use aseptic technique!
1. Add 2mL RDM medium to each well of the 6-well plate.
2. Following the diagram below, add 200 uL of media OR inoculate 200uL of each bacterial
strain to duplicate wells. Mix well by pipetting gently.
3. Carefully place your 6 well plate in the Ziploc bag without disturbing the media, and tuck
the open end of the bag under the plate (do not seal the bag).
4. Label half of the LB plate with “wild-type” and the other half with “mutant”. Using the
overnight broth cultures, streak the appropriate strain in each sector to obtain single
colonies.
5. Incubate both the 6-well plate and LB plate at 30 C for 2 days.
6. After 2 days, carefully remove both plates from the incubator.
For all of the following steps, be careful to pipet in and out from the same location on each
well (i.e. the bottom) to avoid disturbing the biofilm and avoid splashing on the walls of the
wells.
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7. From the 6-well plate, remove the liquid portion of the biofilm using a pipette. Avoid
disturbing the biofilm.
8. Stain each well with 2mL of filtered crystal violet, and let sit for 20 minutes.
9. Remove crystal violet using a pipette.
10. Add 2mL of distilled water to each well, then remove carefully using a pipette. Each
well should be rinsed a total of 3 times to remove excess crystal violet.
11. Add 2mL of 95% ethanol to each well and let sit for 5 minutes. The ethanol will help
dissolve the biofilm.
12. Pipette up and down to mix this suspension as best you can, scraping to remove biofilms
stuck to the sides or bottom of the wells.
13. From each well, remove 0.3mL of this suspension and place in a spec tube containing 2.7
mL of water. Mix well by pipetting. Read the OD at 570nm. Construct a data table
showing the A570 for each well of your 6-well plate (take dilution factors into account).
14. Observe the bacterial growth on the LB plate and record your findings. If single colonies
are not yet visible, incubate the plate for 2 more days at 30 C or for up to 7 more days at
room temperature.
Post-lab Questions
1. Which strain is better at forming a biofilm, wild-type or mutant? Describe the evidence you
have to support your conclusion.
2. What was the purpose of inoculating two wells with media only? How did the A570 for these
wells compare to the A570 to the other wells? How can the A570 values from the media only
wells be used to help you compare the biofilm-forming ability of the wild-type and mutant
strains more accurately?
3. Of the six A570 values that you obtained, which ones are expected to be identical to each
other? Were they identical to each other? If not, explain possible reasons why not (be as
specific as possible).
4. On the LB plate, did both strains appear identical? If not, specify how they looked different.
5. Considering your observations from the LB streak plate and 6-well plate, form a hypothesis
for why one of the strains is better at forming biofilms than the other strain.
99
AMES TEST FOR MUTAGENICITY
OBJECTIVES: Students should be able to
1. Explain the biological basis of Ames test and its limitations
2. Perform an Ames test and interpret the results
Cancer is one of the major causes of morbidity (illness) and mortality (death) in America. While
all the factors that contribute to its onset are not known, it is clear that there are chemical and
physical agents that can induce cancer. These agents are called carcinogens. If carcinogens can
be identified, then it may be possible to reduce their levels in the environment. The most
definitive way to detect carcinogens is to inoculate a sample into animals and monitor for the
development of tumors. This process is expensive, time consuming, and cumbersome. Imagine
having to perform such experiments on all new chemicals. Therefore a rapid, economical
screening method is used to distinguish between compounds that might be carcinogenic and
those that are likely to prove harmless. The more demanding tests can then be performed on a
limited number of materials.
The Ames test is a screening assay for carcinogens that uses bacteria to detect chemical
mutagens. It is based on the premise that most carcinogens induce cancer because they are
mutagens. If a substance is shown to be mutagenic for bacteria, it may also alter DNA in
eukaryotic cells, and more tests would be needed to determine whether the substance is a
carcinogen.
Recall that an auxotroph is a bacterial mutant that cannot synthesize one or more of its amino
acids, so it can only grow if provided with the amino acid it cannot make for itself. Conversely,
a prototroph is a bacterium that can synthesize all of its own amino acids, so it can grow in a
minimal medium that is not supplemented with amino acids. In the Ames test, a special triple
mutant strain of Salmonella typhimurium is used. The Ames strain is auxotrophic for histidine
(his-), is defective in repair of new mutations (uvrB-), and has a defective outer membrane (rfa-).
The rate of reversion (back mutation) of the his- mutation to prototrophy (his+) caused by the
chemical is measured in the Ames test.
The test is performed by first spreading the Salmonella onto a minimal medium plate. Then, the
suspected mutagen is placed on a disk on the plate so the chemical can diffuse out of the disk
into the media. The minimal medium used in the Ames test contains a very small amount of
histidine that supports only a few rounds of bacterial cell division. This small amount of
histidine is essential because many mutagens work only on replicating DNA. If a back mutation
to prototrophy occurs, visible colonies develop. The reversion rate is compared to a control plate
(no chemicals added) and is proportional to the mutagenicity of the chemical.
Some versions of the Ames test include an extract of mammalian liver enzymes in the media,
since some chemicals only become mutagenic and carcinogenic after they have been metabolized
100
in the liver. In this lab, you will perform an Ames test without liver enzymes.
MATERIALS (perform the Ames test with your lab group)
•
•
•
•
•
•
•
•
Glucose minimal-salts agar plates [glucose minimal-salts, 0.05 mM histidine and 0.05
mM biotin]
Complete medium plate
Sterile filter paper disks
Micropipettor and sterile tips
Forceps and alcohol
Sterile tubes (for dissolving solid test chemicals)
Test chemicals:
• Sterile Water (negative control)
• 2-nitrophenylenediamine 100 µM (positive control)
• A household item of your choice: (ex.: cosmetic item, hot dog, cigarette, hair dye)
**If you bring in a food item, it cannot be consumed after the lab. Dangerous
items that could harm others (such as pepper spray) cannot be used.** You may
want to bring a back-up test item in case the TA does not approve your first item.
Culture
• Salmonella typhimurium his- uvrB-, rfa- (Ames no. TA-1538) in TSB
PROCEDURE
Salmonella typhimurium is a potential pathogen. Be certain that all materials used in this
experiment are disposed of properly as indicated by your instructor. The chemicals may be
mutagenic; handle with care.
1. Solid test chemicals should be dissolved in a sterile tube with a small amount of sterile
water. (not necessary for liquid test chemicals)
2. Label your minimal medium and complete medium plates. For each plate, pipette 100
µL of the Salmonella and perform a spread plate to spread an even layer of the bacteria
over the entire surface of the plate.
3. Using flame-sterilized forceps to prevent contamination, dip a filter disk into the test
chemical solution, blot off the excess, and place on one half of the test plate. On the other
half of the test plate, use a filter disk dipped into a more dilute solution of the test
chemical (record the dilution factor used). Label each half of the plate.
4. Control plates: Dip a filter disk into sterile water and place on one half of the control
plate (negative control). Dip a filter disk into 2-nitrophenylenediamine and place on the
other half of the control plate (positive control).
5. Ensure that all excess liquid has been absorbed into the plates before placing all plates
into the 37°C incubator for 48 hours.
101
6. Count the number of colonies and describe their location.
Post-lab Questions
1. Why must control plates be included in the assay each time it is performed?
2. Is the pattern of growth different on the minimal plate vs. the complete medium plate?
Explain how and why you expect the pattern of growth to differ on both plates.
3. Why does the test use a bacterium that is defective in outer membrane synthesis and
DNA repair?
4. What advantage is there to starting with an auxotroph and measuring reversion to
prototrophy rather than starting with a prototroph and measuring mutation to auxotrophy?
5. Why aren't all carcinogens mutagens?
102
PLAQUE COUNTS
OBJECTIVES: Students should be able to
1. Explain the biological basis for plaque formation
2. Enumerate the virus titer in a suspension
3. Calculate virus populations from PFUs on plates
4. Demonstrate the soft agar overlay method
Viruses are noncellular obligate intracellular parasites. They are composed of nucleic acid
surrounded by a protective protein coat (and in some cases a phospholipid envelope). Viruses
are metabolically inert outside of a host cell. They can only replicate using the machinery
provided by the living host. Viruses are replicated in the laboratory using cells of the appropriate
host. Animal viruses are grown in tissue culture, on a monolayer or a suspension of animal cells.
Bacterial viruses, or bacteriophage, are grown in soft agar or in broth seeded with bacterial
cells.
Phage replication involves five main steps: Attachment or adsorption, Penetration, synthesis,
Assembly, and Release. Lytic viruses lyse the cell during the release step of the phage
replication cycle. Thus, we can quantify the concentration of phages by making dilutions of the
virus and assaying on a seeded agar surface for plaque formation. Plaques are clear areas on a
lawn of host cells that represent the point at which a single infectious virus particle was
deposited. As a result of subsequent lytic cycles, the host cells in the region are destroyed,
resulting in an area of clearing. The size of the plaque is dependent on the replication time of the
virus and the generation time of its bacterial host.
A plaque forming unit (PFU) is analogous to a CFU of bacteria: a plaque represents where a
single infectious virus particle was seeded in soft agar with bacteria, whereas a colony represents
where a single viable bacterium was originally deposited on an agar plate. A bacteriophage
plaque count can be used to determine the titer, the concentration of phage particles, in a
suspension of phage.
𝐱 TDF
Original Concentration (PFU/mL) = # of PFU
Volume of phage used (ml)
MATERIALS
•
•
•
•
Soft Agar
TSA
Broth culture of Escherichia coli B
Suspension of T4 Phage
103
PROCEDURE
Note: Do not remove soft agar from water bath until you are certain you are ready to proceed
with that step. The soft agar will solidify quickly at room temperature.
1. Pour 3 TSA plates. Allow them to solidify before continuing.
2. Dilute T4 phage suspension to total dilution factors of 10, 100, 1000, and 10000 with
TSB.
3. Working quickly, add 250 ul of E. coli B to three different soft agar tubes. Swirl
gently to mix.
4. Add 0.1 ml of phage from the 100, 1000, and 10000 dilutions to the soft agar tubes.
Swirl gently to mix well.
5. Pour soft agar containing E. coli and T4 phage dilutions onto separate TSA plates,
swirl plate to evenly distribute agar. Allow it to solidify.
6. Incubate at 37C and calculate PFU concentration of original phage suspension during
the next class.
Post-lab Questions
1. Why do plaques stop increasing in size?
2. A virus suspension is enumerated by counting particles in the electron microscope and by
a plaque assay. By which method would you expect a higher count? Explain.
3. Bacterial colonies may be observed growing in the plaque. Why would these develop?
4. How would you perform a plaque assay for lytic animal viruses?
5. How would you determine the total bacteriophage population in your environmental
sample?
104
KIRBY-BAUER: ANTIBIOTIC SUSCEPTIBILITY TESTING
OBJECTIVES: Students should be able to
1. Explain the purpose of the Kirby-Bauer test.
2. Describe how the Kirby-Bauer test works, how its results are interpreted, and what are
limitations of the test.
3. Describe the modes of action of the antibiotics used in the experiment.
Antibiotics have helped control disease and saved countless lives. Five main modes of action of
antibiotics are:
1. Inhibiting cell wall synthesis (ex: Ampicillin, Penicillin G)
2. Inhibiting protein synthesis (ex: Erythromycin, Chloramphenicol, Streptomycin,
Tetracycline)
3. Inhibiting nucleic acid synthesis (ex: quinolones, Rifampin)
4. Disrupting cell membranes (ex: Polymyxin)
5. Acting as an antimetabolite (ex: Sulfamethoxazole-trimethoprim)
To be effective, the appropriate drug must be used, and its choice is determined by several
factors including the site of infection, the tolerance of the host and the nature of the pathogen.
Drugs are usually administered prior to isolation of the infectious agent, and often diagnosis is
confirmed without isolation as well. In cases where the pathogen is isolated, drug sensitivity
may be determined.
There are several methods that can be used to measure antibiotic susceptibility. Among them is
the disk diffusion method in a procedure commonly called the Kirby-Bauer technique.
Antibiotic impregnated disks are placed on the surface of a solid medium (typically the rich
medium Mueller-Hinton agar poured to a uniform thickness) across which has been spread a
known concentration of the isolated pathogen. After 24 hours incubation, each antibiotic has
diffused from the disk into the agar. Antibiotics that inhibit microbial growth produce a clear
zone around the disk in which no organisms grow. The diameter of this zone of inhibition is
compared to a standardized chart that indicates whether the bacterium is sensitive, intermediate
or resistant (SIR) to the drug. A bacterial strain may be categorized as “resistant” even if a zone
of inhibition is present. The SIR chart designations have been determined by the drug
manufacturer from many experiments that compared zone size to the drug concentration attained
in the body at the site of infection. Sensitive designations indicate that the drug will inhibit the
organism in the infected person. If the microbe is considered resistant, then the drug will be of
no use clinically. The intermediate designation suggests that the drug might be useful if there are
no better alternatives.
This test measures susceptibility to many different drugs in a single, easily performed procedure.
It cannot be used, however, for bacteria that are slow growers or too fastidious to grow on the
standard assay medium.
105
MATERIALS – Work in groups of 4.
•
•
•
•
•
Forceps
Sterile swab
Mueller Hinton agar in large 150 mm petri plate
Cultures (broth):
• Escherichia coli
• Staphylococcus aureus
• Pseudomonas putida
Antibiotic Disks:
• Ampicillin (10 mcg)
• Erythromycin (15 mcg)
• Chloramphenicol (30 mcg)
• Penicillin G (10 units)
• Streptomycin (10 mcg)
• Sulfamethoxazole-trimethoprim (300 mcg)
• Tetracycline (30 mcg)
PROCEDURE
1. Prepare a spread plate by inoculating 0.5 ml of the culture onto the plate; spread using the
swab as you rotate the plate so that the entire surface is evenly coated with the culture.
2. Wait for a few minutes until all excess liquid is absorbed in to the agar.
3. Dispense the disks – note the drug designation and the potency of each drug used. IF
THE DISK SHOULD FALL ONTO AN INCORRECT AREA OF THE PLATE,
QUICKLY MOVE IT. GENTLY TAP THE DISK INTO PLACE.
4. Incubate the plate right side up (e.g. agar side down) at 37oC for 24 hours for E. coli and
S. aureus. P. putida needs to be incubated at 30 oC for 24 hours.
5. Measure the diameter of the zones of inhibition across the disk to the nearest millimeter.
If there is no visible zone of inhibition, record the diameter as 0.
6. Record your results and determine the interpretation as S (sensitive), I (intermediate) or
R (resistant) by comparing to the SIR chart (next page). Make sure that you record
results for all the organisms.
Post-lab Questions
1. Based on these results, which drug could be used to treat S. aureus? E. coli? P. putida?
2. What factors might influence the size of the zone of inhibition?
106
3. What is the mode of action of each drug? Which drugs are more effective against gram
positive bacteria? What other factors must be considered when selecting an antibiotic for
treatment?
4. Explain why a bacterium might be considered resistant to a drug even if you observe a
zone of inhibition.
5. Describe two mechanisms by which a bacterium can develop resistance to an antibiotic.
6. See the research article at https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7758518/.
This study tested the antibacterial effects of water extracts or methanol extracts of
Jacaranda flowers using the Kirby Bauer disk diffusion assay. Which extracts had
stronger antibacterial effects?
107
SIR INTERPRETIVE CHART
ANTIMICROBIAL
AGENT (amount per disc)
ZONE OF INHIBITION (mm)
RESISTANT (R)
INTERMEDIATE
(I)
SUSCEPTIBLE
(S)
AMPICILLIN (10μg)
E.coli
S.aureus
P.putida
≤13
≤28
≤11
14-16
--12-14
≥17
≥29
≥15
CHLORAMPHENICOL
(30μg)
E.coli
S.aureus
P.putida
≤12
≤12
≤12
13-17
13-17
13-17
≥18
≥18
≥18
PENICILLIN (10U)
E.coli
S.aureus
P.putida
≤14
≤28
≤14
----------
≥15
≥29
≥15
11-15
11-15
11-15
≥16
≥16
≥16
≤14
≤14
≤14
15-18
15-18
15-18
≥19
≥19
≥19
STREPTOMYCIN (10 μg)
< 11
12-14
>15
ERYTHROMYCIN (15
μg)
<13
14-22
>23
SULFAMETHOXAZOLETRIMETHOPRIM (300
≤10
μg)
≤10
E.coli
≤10
S.aureus
P.putida
TETRACYCLINE (30μg)
E.coli
S.aureus
P.putida
108
EPIDEMIOLOGY: A SIMULATED EPIDEMIC
(Lab adapted with permission from P. Fidopiastis, Cal Poly SLO Microbiology)
OBJECTIVES: Students should be able to
1. Define “epidemiology”.
2. Describe the contributions of Dr. John Snow to modern epidemiology.
3. Calculate and describe the significance of Prevalence, Incidence, and Contact-Specific
Attack Rate.
Read the following two pages before today’s lab:
https://www.cdc.gov/csels/dsepd/ss1978/lesson1/section1.html
https://www.cdc.gov/csels/dsepd/ss1978/lesson1/section2.html
Epidemiology is the study of public health and its applications. Epidemiologists study the
distribution of health events, focusing on their frequency in the population and their pattern
(ie: Who is affected? When? Where?). Epidemiologists also study the determinants or causes
influencing health events.
The first modern, systematic, scientific epidemiologic study was conducted in 1854 by John
Snow, a London physician who recorded and mapped the locations of cholera cases in the city
during an outbreak. He examined death records and made a spot map of locations of death in
London to try to determine the source of the outbreak. At this time, the causative bacterial agent
of cholera (Vibrio cholera) and the source of infection (contaminated water or food) were not yet
known. During the first week of September alone, 600 people died! After carefully plotting the
cases on a map and analyzing the data, Dr. Snow found that cases were centered around the
Broad Street pump, which supplied water to the nearby area. The water intake for that pump was
from a polluted region of the Thames River. He tracked the source of the epidemic to
contamination of the intake region by sewage. To test whether this particular pump was one
source of the outbreak, the pump handle was removed so that water could not be collected from
that location. After the pump handle was removed, the cholera outbreak subsided, indicating that
contaminated water from this source was at least one cause of the cholera outbreak in the city.
The shape of an epidemic curve helps to distinguish the likely origin of an epidemic. To
generate an epidemic curve, start by plotting the index case and then the number of new cases
during a specified time period (such as number of cases per day). During a common (point)
source epidemic (i.e. caused by contaminated food or water), the epidemic curve is characterized
by a sharp rise to a peak in time, with a rapid decline that is typically less abrupt than the rise.
Cases continue to be reported for about one incubation period for the disease. In contrast, during
a propagated (host-to-host) epidemic, the epidemic curve is characterized by a relatively slow,
progressive rise. Cases continue to be reported over several incubation periods of the disease. In
the case of vector-borne diseases, the case distribution often has apparent spatial and temporal
components to that are related to vector (i.e. arthropod) life cycles.
109
Today you will be approximating the work of an epidemiologist. Each of you will be exposed to
a potential “pathogen” via a handshake. Only some individuals in the class will initially contact
the pathogen. Those that are infected will spread the pathogen to others in the class by
subsequent handshakes.
Epidemiology Data Analysis
Point prevalence is the proportion of the population with the disease at a specific time point.
Point prevalance does not give any information about when the disease developed or how it was
caused.
Point Prevalence (%) = Total # of cases (new and preexisting) at a specified point in time x 100
Total # of individuals in the population at the same point in time
Incidence proportion (attack rate or risk) is the proportion of a population that develops
disease during a given time period and is calculated by the following:
Incidence proportion =
# of new cases occurring in a population during a specified time interval x 100
# of individuals in population at risk at start of time interval
Point prevalence is a slice through a population at a point in time to determine who has the
disease but does not give information about when the disease developed. Incidence proportion
measures new cases of disease in a population and thus measures risk – the denominator
indicates the number of individuals that may potentially develop disease.
Contact-specific attack rate is a subtype of incidence proportion. The denominator includes all
susceptible individuals that were exposed to the agent (a person or a contaminated food item).
Time may not be explicitly defined, but the incubation period is usually determined. The
contact-specific attack rate may be specified for a given exposure, i.e. calculated for each type of
food eaten.
Contact-Specific Attack rate for simulated epidemic lab exercise =
# of people who shook hands with a particular individual and became infected
Total # of people who shook hands with that individual
Example calculation:
Swab #
1
2
3
4
5
# Infected
2
2
3
1
0
# Not Infected
1
1
0
2
3
Total
3
3
3
3
3
110
Contact Specific Attack Rate (%)
67
67
100
33
0
PROCEDURE
1. Work individually.
2. On the bottom of one SD plate (Synthetic dextrose minimal medium for yeast) draw and label
3 sectors: #1, #2, and #3, as illustrated in the Worksheet below.
4. Record the number of the swab placed at your desk on your worksheet. Some swabs
have been dipped in a culture of Saccharomyces cerevisiae, and others have been dipped in
sterile TSB.
5. Stop at this point. Wait until the instructor gives you the signal to begin the experiment.
6. When instructed, place a glove on the hand you do not write with (to be used for hand
shaking). Do not let the glove touch any surface. Generously rub the swab over the gloved hand
– cover every portion of the hand, both the front and back. Place the swab in the autoclave
waste for disposal.
7. Stop. Wait for the instructor’s signal.
8. When instructed, shake the gloved hand of one person from the other side of the room. Only
shake hands once – do not shake hands with more than one person during each turn.
Record on your worksheet the name and swab number of the student you shook hands with under
“Handshake #1”.
9. Without touching the glove to any other surface, inoculate section #1 of your SD plate by
rolling the tips of your fingers (especially the thumb) across just section #1.
10. Stop. Wait for the instructor’s signal.
11. When instructed, shake the gloved hand of a second student in the class. Do not shake
hands with a person you have previously shaken hands with. Only shake hands with one
person. Record on your worksheet the name and swab number of the student you shook hands
with under “Handshake #2”.
12. Without touching the glove to any other surface, inoculate section #2 of your SD plate by
rolling the tips of your fingers (especially the thumb) across just section #2.
13. Stop. Wait for the instructor’s signal.
14. When instructed, shake the hand of a third student in the class. Do not shake hands with a
person you have previously shaken hands with. Only shake hands with one person. Record
on your worksheet the name and swab number of the student you shook hands with under
“Handshake #3”.
111
15. Without touching the glove to any other surface, inoculate section #3 of your SD plate by
rolling the tips of your fingers (especially the thumb) across just section #3. Discard glove in the
autoclave waste.
16. On each of the 3 sections of the completed SD plate, write the number of the person’s swab
you inoculated on that section below the section number.
17. Place the plate carefully labeled with your full name in the 30°C incubator until next time.
Next lab:
1. Examine each sector of your plate for colonies of S. cerevisiae. If you are curious, you can
confirm colonies are S. cerevisiae and not a contaminant by making a Gram stain and looking for
large budding cells typical of yeast.
2. Record your results from each sector on the data sheet (Table 1) under “Individual data”.
3. Report your data to the class, on the board or computer. Once you have all the data,
calculate your specific attack rate for your swab number and record this in the class data
as well. Once completed, copy the final data onto the “Class Data” area of the worksheet.
4. Working in your groups, calculate the following using the class data: point prevalence (after
the first, second, and third handshakes), incidence proportion (during the second the third
handshake periods), and contact specific attack rates.
5. How many carriers of S. cerevisiae resulted after Round 1? After Round 2? After Round 3?
6. As a group, try to figure out who the original carrier(s) of the infectious agent were. Start by
looking at the contact specific attack rate for each person. Also look at the first handshake. Did
both people who shook hands with each other indicate infection on their first sector? Did the
suspected individual(s) transmit the pathogen during each subsequent handshake? Also look at
the outcome for handshakes from each infected individual. Does the suspected carrier infect an
individual with each subsequent handshake?
7. Make sure that all of the class data are included and discussed in your lab notebook.
112
Worksheet: Fill this out carefully as you shake hands.
Rules: Do not shake hands until the instructor signals to do so during each of the 3
handshakes. Shake hands with only one person during each turn. Shake hands with a
different person during each turn.
Your swab # ____________
CALCULATIONS:
POINT PREVALENCE (3 different calculations: after first handshake, after second
handshake, and after third handshake):
INCIDENCE PROPORTION (2 different calculations: during second handshake period
and during third handshake period):
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TABLE. EPIDEMIOLOGY DATA ANALYSIS: TRANSMISSION OF S. CEREVISIAE
Individual data from the simulated epidemic
Agent Present or Absent? (S. cerevisiae
Plate sector number
Swab number
on plate?)
1
2
3
Class Data from the simulated epidemic
Swab
number
1st sector swab #
(circle if + for S.
cerevisiae)
2nd sector swab
# (circle if +)
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
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3rd sector swab
# (circle if +)
Contact Specific
Attack Rate (%)
TERMITE GUT SYMBIONTS
OBJECTIVES: Students should be able to
1. Observe microbes from all 3 domains in the termite hindgut.
2. Explain the important roles of the different types of microbes found in the termite hindgut.
Many microbes live in symbioses in nature. Termites can feed on cellulose (a β(1->4) linked Dglucose polysaccharide that is a major component of wood and paper) only because the termite
hindgut is full of eukaryotic, bacterial, and archaeal microbial symbionts that can break
down cellulose into compounds that the termites can digest. It is estimated that 90% of the
biodiversity in the termite hindgut cannot be cultured. The termite gut is microoxic and anoxic,
but rich in hydrogen gas (see diagram below from Brune and Friedrich, Current Opinion
Microbiol, 2000). In this lab, you will have an opportunity to view the diversity of microbes
present in the termite gut.
The intestinal flagellates that we can easily see are
likely the major source of hydrogen in the gut. These
gut protozoa are often associated with methanogens
(archaea producing methane from hydrogen and
carbon dioxide) and acetogens (producing acetate
from hydrogen and carbon dioxide) that may be
endosymbionts of the protozoa or associated with the
protozoan surface.
Reactions that the microbes perform in the termite gut
include hydrolysis of cellulose and hemicellulose
(hemicellulose is related to cellulose but has shorter
polysaccharide chains and other sugars besides
glucose), fermentation of the hydrolysis products to
short chain fatty acids that are used by the host, and
nitrogen cycling and nitrogen fixation. Bacteria and
Archaea are present in the hindgut of termites at high
concentrations – 107 – 1011 per ml.
Methane production in termites contributes
significantly to global methane production! Methane
is a potent greenhouse gas. Methanogens can be
identified by the autofluorescence of a unique
coenzyme – F420. Coenzyme F420 plays an
important role in the metabolism of C1 compounds in
methanogens. F420 fluoresces under UV light.
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From Brune, A., Nature Reviews Microbiology, 2014
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PROCEDURE (1 termite per group)
1. View minutes 5-6 of this video: http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2556161/ for
the dissection procedure. Termites are provided on ice. Keep the termites on ice in a plastic petri
dish before and during dissection.
2. For optimal viewing, transfer the hindgut to the microscope slide while minimizing waiting
time and exposure of the sample to the air. If termites are small, you may transfer the entire gut
to the slide. If termites are large, use forceps to only transfer a small portion of the gut (no larger
than the width of cover slip) to the slide.
3. Once the hindgut has been transferred to the microscope slide, add a drop of water to the
sample to ensure no air is present when the cover slip is applied. Alternatively, a cover slip can
be placed over the hindgut sample and water can be pipetted to fill in any air gaps. Remove any
excess water with paper towel or Kimwipe.
4. Seal the perimeter of the cover slip with clear nail polish to fix the sample to the slide.
5. Observe the termite hindgut using both your microscope (brightfield microscopy) and
fluorescence microscopy.
Post-lab Questions
1.
2.
3.
4.
What are the electron acceptor and donor for methanogenesis?
In which domain are the organisms that perform methanogenesis found?
Spirochetes can be abundant in the termite gut. How can you identify them?
Do you expect any of the microbes in the termite gut to have nuclei? Explain.
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EUKARYOTIC MICROBES
OBJECTIVES: Students should be able to
1. Describe the various categories of organisms that are eukaryotic microbes.
2. Contrast their observations of eukaryotic microbes with prokaryotic microbes.
Eukaryotic microbes are a very diverse and beautiful group of organisms. There are various
ways to classify eukaryotic microbes, but in general the two main groups of eukaryotic microbes
are fungi (such as yeast and mold) and protists (including protozoa and microscopic algae).
Protists can be further classified based on how they move: (1) flagellates move using flagella;
(2) amoeboids move using pseudopodia; (3) ciliates move using cilia; and (4) apicomplexa are
non-motile and have a special organelle called an apicoplast.
Many eukaryotic microbes have characteristic components in their cell envelopes. Fungi have
cell walls containing glycoproteins. Protozoa do not have cell walls, but instead have a pellicle,
which is a supportive proteinaceous layer underneath the plasma membrane. Microscopic algae
may have cell walls containing cellulose or silica.
In general, eukaryotic microbes are larger than prokaryotes, but eukaryotic microbes can vary
tremendously in size. For example, Saccharomyces cerevisiae (baker’s and brewer’s yeast) is
about 5-10 μm in diameter whereas Paramecium species can be over 100 μm long. Eukaryotic
fungi have a critical environmental role in decomposing organic compounds. Microscopic algae
such as diatoms and dinoflagellates have a major role in performing photosynthesis. Still other
eukaryotic microbes are pathogens, such as Entomoeba histolytica (an amoeboid that causes
dysentery), Giardia (a flagellate that causes hiker’s diarrhea), and Plasmodium species
(apicomplexans that cause malaria).
Here, you will observe yeast, Paramecium (a ciliate), and Euglena (a flagellate). The yeast are
stained with a pH-sensitive dye, Congo Red. At pH>5 Congo Red is red; between pH 3 and 5 it
is purple; and at pH<3 it is blue. Both Paramecium and Euglena are commonly found in
freshwater and saltwater environments. Paramecia are heterotrophs that eat small microbes such
as yeast. They use their cilia to propel themselves through water in a spiral motion. They also
use cilia to move food into the oral groove then the gullet, where food vacuoles form to digest
the food. Euglena can gain its nutrients as a heterotroph (ingesting food via phagocytosis) or as a
photoautotroph (using its chloroplasts to perform photosynthesis). Euglena has a single polar
flagellum and a red eyespot (due to carotenoid pigments) that is believed to aid in phototaxis.
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MATERIALS
•
•
•
•
•
•
•
Paramecium caudatum
Congo red-yeast suspension
Euglena acus
Protoslo or 1.5% Methylcellulose (optional)
5% acetic acid
transfer pipets
coverslips
PROCEDURE
1. Observe Paramecium feeding on Congo red-stained yeast by performing a wet mount. Use a
transfer pipet to place a drop of the Paramecium culture on the slide. Place a drop of yeast
suspension right next to the Paramecium drop. Place one edge of a coverslip into a drop, letting
the fluid run along the coverslip, then gently lower the coverslip (trying to avoid air bubbles).
Observe the Paramecia and yeast, noting their relative sizes. Note the type of locomotion of
Paramecium. Can you observe both red and blue food vacuoles? Usually, feeding on yeast will
slow the Paramecia down enough to allow observation, but if necessary you can add a drop of
Protoslo to slow the Paramecia down even further.
2. On a different slide, perform a wet mount of Euglena by placing a drop on the slide and using
a coverslip. How does its movement, size, and appearance compare to Paramecium and yeast?
Can you see the red eyespot?
3. After observing Euglena, place a small drop of acetic acid at one edge of the coverslip and
allow the acid to diffuse under the Euglena coverslip. How does Euglena respond?
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Post-lab Questions
1. How do the sizes of Paramecium, Yeast, and Euglena compare? Sketch a cell of each one in
your notebook, showing their sizes relative to each other. Also, sketch in an E. coli cell in your
diagram, showing its size relative to the eukaryotic microbes.
2. In Paramecium, food vacuoles form at the end of the gullet, then move through the cytoplasm
as digestion proceeds. When Paramecia are feeding on Congo red-stained yeast, where would
you expect to find red food vacuoles? Where would you expect to find blue food vacuoles?
3. Suppose you could shine a light at one edge of the Euglena slide. How would you expect the
Euglena to respond?
4. Prior to today, all of your microscopy in this course has been with bacteria. How is the
microscope procedure you used to observe eukaryotic microbes different from the procedure you
have used to observe bacteria? (Consider magnification, staining, etc.)
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Second group project plan: Guidelines
1. Brainstorm with your group to formulate a hypothesis or question involving bacteria that you
can obtain from the environment. Be creative – choose a question that you are really curious to
discover the answer to! This is the main part of the second project (about two-thirds to threequarters of the project).
o Design a plan to test your hypothesis.
o At least 1 experimental method used to test your hypothesis should come from the
following protocols: Biofilm, Ames Test, Plaque count, Kirby-Bauer. Choose the most
appropriate test to use, depending on your hypothesis.
o Include any other methods from the course that are needed to address your question.
o A minimum of two separate experiments should be conducted to test your hypothesis.
The protocols for these two experiments should be written out completely, step-by-step
how you will perform the experiment. This will include concentrations, volumes,
dilutions, incubation times and temperatures, any measurements that will be made, as
well as any and all controls.
o Discuss your ideas with your TA.
o Your plan should be logical and thoughtful, and based on the primary literature turned in
with the plan.
2. From your environmental sample, choose at least two isolates that look quite different from
one another. This is a minor part of the second project (about one-third to one-quarter of the
project).
o Formulate a second hypothesis regarding the identity of the bacteria, based on where your
bacterial sample came from.
o With these two isolates, conduct a Gram stain and two tests from Module 1 to see
whether the data from these tests support your hypothesis on the identity of your bacteria.
You do not need to definitively identify the organism in this part of the project, since you
are only doing a Gram stain and two tests; the focus of the second group project is your
first hypothesis (step 1).
3. In general, only bacteria to be used as controls can be requested from the Biol 302L staff.
4. Be creative in designing your first hypothesis (from step 1) to earn the creativity points for the
second group project!
5. Turn in the first page of 4 primary literature articles supporting your hypotheses.
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Biol302(Group(Project(Plan(Schedule
Group&name:&__________________________________________
Section&Number:&_______________________________________
Request&date:
Experiment
Materials
Experiment
Materials
Experiment
Materials
1
Start&date:
2
3
4
5
Request&date:
1
Start&date:
2
3
4
5
Request&date:
1
Start&date:
2
3
4
5
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General Instructions for Power Point Presentation:
How to present original work effectively
This is a guide to help you prepare your second group project presentation. First, consider
your audience (your classmates and TA). Although they have similar background knowledge as
you, unlike you they have not been immersed in your project for the past few weeks.
The essential elements of a good presentation are:
1. Ample but brief Background information
2. Clear and concise Hypotheses and Rationale
3. Description of Methods used (with figures/flow charts if applicable)
4. Clear graphical/tabular presentation of important Results
5. Clear and concise Conclusions
6. Discussion of Future work
Some general guidelines:
o Limit wordiness; your slides should not say everything you will say out loud, but should
only illustrate your main points.
o Include figures and/or diagrams where possible; more visuals and fewer words are always
better. However, make sure these are relevant and not too busy or distracting.
o If you don’t have a lot of figures, that’s ok, but don’t fill space with lots of words.
Specific guidelines: Include the following slides, keeping to the guidelines for content and
length:
1. Title slide: Include the title of your work, the names of the scientists who conducted the
study, and the institution with which you are associated (CSUF). You should include the
department (Biological Science) with which you are associated. [1 slide]
2. Background: Include relevant background to bring your audience up to speed regarding
why you chose this particular study to conduct. What is the big picture? What is the
question you are interested in? Why is this question important? Include background
information on the organisms you are using. Include figures if you can. [2-4 slides]
3. Hypotheses: Clearly and concisely state your hypotheses. The Background in the
previous section should have made clear your rationale leading to these hypotheses. The
rationale may be shown on the slide, but does not need to be stated on the slide if you can
adequately explain it verbally. Have a max of 1 slide per hypothesis. [1-2 slides]
4. Methods: Briefly describe the methods that you used. Be specific enough for your
audience to follow what you have done without giving excessive detail. Describe the
controls and explain how they helped you interpret your results. Do not describe in detail
any methods that were used in class, but explain any modifications you made to those
procedures. Flow charts or diagrams of a set-up are often helpful. There should only be
ONE slide with condensed materials and methods for the bacterial identification part of the
project. [2-3 slides]
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5. Results: Show Results figures and tables and describe what is shown in each figure and
table. In a table or graph, only averages from repeated experiments (not raw data) should
be presented. Make sure your figures and tables are clear enough to be easily understood,
but detailed enough to stand alone. Also, make sure all text is large enough to be seen (for
example, make sure the axis labels on a graph are readable). Try not to put too many
tables/graphs on one slide. Limit your usage of text; use a few bullet points to explain the
results. [2-4 slides]
6. Conclusions: Briefly list the main conclusions you made from your study. Do not restate your results, but rather explain their significance. How do the results relate to the
overall question you were trying to answer? Do your results support or refute your
hypotheses? Why or why not? Explain any sources of error you feel may have influenced
your data, or any data that you threw out and why. [1-2 slides]
7. Future Work: In light of what you have discovered by conducting this study, what
would be your next step(s) if you continued this investigation? What new questions arise
from your data/conclusions? If parts of the study were unsuccessful, what alterations
would you make to the current study to make it more successful? [1 slide]
8. Literature Cited: List any references you mentioned in your presentation. [1 slide]
9. Acknowledgements: List the names of those individuals who helped you along the way
(i.e. provided you with meaningful discussions, helped you analyze your data, or provided
with any outside products, etc.). Verbally mention how they helped you. This section is
optional. As is the case for the paper, you may acknowledge the lab technician but DO
NOT acknowledge your TA. [1 slide].
The presentation should be between 10 to 15 minutes long (followed by 3-5 minutes for
questions). Keep your presentation focused, logical, and clear. Remember your audience!
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Final Group Paper: Instructions for writing and formatting
Title (16pt font – use Times New Roman for the WHOLE paper) should be centered. Exercise
care in composing a title. Avoid the main title/subtitle arrangement, complete sentences, and
unnecessary words. The title is not to exceed 20 words (including indefinite articles and
prepositions such as “a”, “the”, and “of”). The title should be informative and descriptive so that
the scope of the study is understood just from reading the title.
By line (14pt font), single spaced, centered. List the authors alphabetically by last name. The
last author serves as the corresponding author so there should be an asterisk next to that name.
Under your names list the institution where the work was done (CSUF) and the department you
are in (Biological Science).
Example:
Cannon, Ryan, Tanya Ledezma, Aaron Salcido, and April Ulloa*
California State University Fullerton, Department of Biological Science
At the bottom of the first page should be a footnote with:
*Corresponding author: full name
E-mail: email address
Abstract (12 pt font). Max of 250 words, single spaced, centered. Make sure that it includes all
the important elements listed below. At the end, indicate the word count in parenthesis ex. (248
words)
• Avoid abbreviations and references, and do not include diagrams. Abstracts can be
published separately from the rest of the paper, so ensure that it is complete and
understandable on its own.
• The abstract should include:
o brief description of the background of your proposed research (but don’t just copy
sentences directly from the background section verbatim).
o the hypotheses of the study
o brief mention of the methods used
o key results and general conclusions
Hint: write the abstract LAST after writing the rest of the paper, since the abstract summarizes
the entire paper. Start with a summary that includes everything you think is important and
gradually cut it down to 250 words by eliminating unnecessary words/information.
Body of the Paper (12 pt. font)
Introduction Max of 2 pages double spaced.
• Give enough background and significance information for your reader to understand and
appreciate the research you will be describing. Provide the context of what was known
prior to the experiments you are about to describe. There should be BOTH (1) some
general background of the field pertaining to the research, and (2) more specific
background relating to your research. The introduction should also address the
importance of your hypothesis and how it will be tested; how may your research expand
on the knowledge of this field, and how will you carry it out?
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• At minimum, your introduction should address all of the following questions.
a. What problem do you hope to address in this field of research?
b. What has already been done in the field? What microbes are usually found in
the environment being investigated and how are they relevant to your research?
Provide two references minimum.
c. What research has already been done that supports your hypotheses? Provide
two additional references minimum.
d. How does the background lead to your hypotheses or research question?
e. What are your hypotheses? State them explicitly in measurable terms.
g. Briefly mention the methods used to test your two hypotheses. (2 sentences
max)
f. What is the significance of your research? How would your hypothesis help
expand on the topic being studied?
• This Background section contains the “literature review” section of your proposal. At
least 4 primary literature articles should be cited here. Reviews are also OK to cite. Use
ASM formatting for in-text citations. NEVER use direct quotes and always
paraphrase/summarize the other studies’ ideas/findings. Remember: Do not plagiarize!
• This is an important place to demonstrate your thoughtful reading and understanding of
the literature, and how it relates to your research.
• The introduction is crucial in framing the rest of your paper; you want to tell a
compelling story with a logical flow
Methods Using past tense and narrative form, briefly describe the methods you used. Procedures
carried out from the lab manual must be described exactly as how they were ACTUALLY
performed (ex. the bacterial culture was diluted by a factor of 5 before creating a smear. The
smear was then air-dried for 2 minutes…). Be specific enough for your reader to replicate what
you did without giving excessive detail. Flow charts or diagrams of a set-up are often helpful.
• Use the power of peer review. Hand your methods to a group member and have them try
to replicate what you wrote. Can they do it exactly as you did, without your help?
Format your methods section as follows:
General: List the names of your materials. Bacteria used, where they were acquired, and
their growth conditions. Any products (toothpaste, mouthwash, antibiotics), etc. that were
used and where they were acquired from (which store, which person, which lab supplied
them to you) or how they were prepared (be sure to mention any sterilization methods
utilized).
Name of Method Used: Once sentence description of why this method was used.
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Description of the general steps that were done, and anything specific that someone who
wanted to repeat your experiment would need to know. Include concentrations and
volumes used, incubation times and temperatures, etc. Be sure to include the positive and
negative controls used with a brief mention as to why these were chosen. (e.x. our positive
control for bactericidal effects was a disk infused with the antibiotic Amikacin. This is an
antibiotic with broad spectrum activity and should readily kill off any organisms near it.) If
your controls don’t give you their expected results, what caused that to happen? (Don’t
write it off as error in the methods too quickly, you may have observed something
interesting).
Name of Second Method Used: Follow the format for the first method used. Describe each
method used in its own section.
Results Max 3 pages, double-spaced, with figures and tables. Include figure captions that
describe what a figure or table is showing; showing the figure or table alone is not sufficient.
(Example text: Based on the findings of Smith et al (2020), garlic was expected to be toxic to
bacteria and inhibit cell growth. The toxicity of the garlic water on strain SYR1 was determined
by a disk diffusion assay. As shown in Figure 1, garlic water caused an average 23±4 mm zone
of inhibition.). Experimental data should be compared to the controls. Mention in text the
statistical significance of the data as well as the general trends observed.
Divide your results section into a minimum of 4 parts with subheadings:
• Isolation of bacteria from [insert environment] – briefly describe the source of the
isolate and why, what the isolates looked like on the plates used (ex. Smooth yellow
round colonies) and give your isolates names that can be used throughout the paper
(ex. SYR1).
• Characterization of Isolates – Describe the tests performed, why they were chosen
to test your hypothesis, and the results of the tests for the isolates named above.
• A subheading to summarize your first experiment to test Hypothesis 1. (Ex.
Garlic inhibited growth of SYR1 on MM, but not TSA.)
• A subheading to summarize your second experiment to test Hypothesis 1. (Ex.
Garlic had no effect on biofilm formation of SYR1.)
Figures/Tables formatting
• The figure or table should highlight what you describe in your results.
• Tables: include a concise legend, informative headings, and units (where appropriate).
• Figures: include a concise descriptive title, an appropriately placed legend, properly
labeled axis with units, a legible font size, and distinguishable data points, lines, or bars.
• In a table or graph, only averages from repeated experiments (not raw data) should be
presented. Make sure your figures and tables are clear enough to be easily understood.
• Figure/table captions should summarize what is being shown in no more than a sentence.
Proceed to describe the most important information being displayed (e.g., What do you
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•
•
•
want your reader to walk away knowing, after looking at this figure/table?)
If appropriate, remember to include 95% confidence intervals on graphs and Standard
Deviations in tables.
All tables and graphs should indicate if the results shown are average values and the
number of samples averaged (Example: Values shown are the average of two separate
experiments. Standard deviation is shown in parentheses).
Pictures are allowed but only include them if they are essential to describing your results.
Any images should be clear enough to easily read and demonstrate the point of the
figure in the printed copy.
Discussion Max 3 pages, double-spaced. The main purpose of this section is to discuss the
significance of your results after briefly summarizing the results. The discussion should address
the following:
• Why are your results important?
• What do they tell you about what you are studying (e.g. what new information have you
learned about biofilms, antibiotics, bacteria, etc.)?
• Compare your results to results in published studies and cite a minimum of two
references here (these can be from references already cited in the introduction).
• Did your results support your hypothesis or not?
• If not, what alternate hypothesis is supported by your data or what additional experiments
would be needed to adequately test your hypothesis?
• Discuss sources of error encountered in your experiments and provide
alternatives/solutions to avoid these sources of error. (If a control gave an unexpected
result, why might this have happened? Don’t write it off as error too quickly)
• Discuss future work; if you were to continue your investigation, what tests would you do
next and why?
Acknowledgements Max 1 paragraph, single-spaced. Acknowledge anyone outside your group
that helped you with your study. If you discussed results with other
groups/classmates, with your family, if someone outside your group bought the toothpaste for
your study, etc. list their names and how they contributed. This section is optional but should be
included if applicable. DO NOT acknowledge your TA! It is your TA’s job to help you, so the
TA does not count as an outside source of help and does not need to be thanked in the
acknowledgements. If you want to acknowledge the lab technician (who prepares and provides
you with all the materials), you may.
References (12 pt font). Your sources should be listed in ASM format. For proper formatting of
references, use the Journal of Bacteriology instructions for authors which can be found at
https://journals.asm.org/references. This resource provides examples for all types of references.
SIX PRIMARY LITERATURE SOURCES MINIMUM.
Grammar. Past tense and third person should be used throughout the paper.
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Instructions for turning in the final group paper:
Submit one copy PER GROUP of the final proposal to the turnitin link in your lab section’s
Canvas site by the deadline specified in the syllabus. Only one member per group should submit
the paper. If more than one paper per group needs to be submitted, it should be approved by the
TA first and will only be granted under special circumstances (i.e. technical problems). A
hardcopy of the final paper must also be submitted by the deadline specified in the syllabus
to your TA’s mailbox in the biology department office and must include the 4 peer reviews
attached.
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