General Microbiology Lab Manual MB 303 - Department of Microbiology [PDF]

General Microbiology Lab Manual MB 303 Fall 2017

©OSU Department of Microbiology, 2010

Instructors:

Dr. Linda Bruslind - Monday & Tuesday lab Dr. Walt Ream - Thursday lab

Note: All MB 303 labs start Week 1, with a no-show drop policy for the class. If you don't attend lab Week 1 you will be dropped from MB 303. ---------------------------------------------------------------------------------------------------Monday section - starts on Monday, Sept. 25, 1:00 pm. Tuesday section - starts on Tuesday, Sept. 26, 1:00 pm. Thursday section - starts on Thursday, Sept. 28, 1:00 pm.

TABLE OF CONTENTS

Laboratory Schedule ..................................................................................................................................... ii Learner Outcomes/Expectations .................................................................................................................. iii Basic Information......................................................................................................................................... iv Course Policies.............................................................................................................................................. v Lab Notebook .............................................................................................................................................. vi Safety Rules ............................................................................................................................................... vii Exercise 1 – Use of the Microscope.............................................................................................................. 1 Exercise 2 – Simple Stains: Microbial Morphology ..................................................................................... 9 Exercise 3 – Transfer of Bacteria................................................................................................................ 11 Exercise 4 – Environmental Sampling ........................................................................................................ 15 Exercise 5 – Differential Stains .................................................................................................................. 19 Exercise 6 – Pure Cultures: Streak Plates ................................................................................................... 23 Exercise 7 – Quantification of Bacteria ...................................................................................................... 27 Exercise 8 – Scientific Writing ................................................................................................................... 33 Exercise 9 – Culturing the Unculturable ..................................................................................................... 31 Exercise 10 – Culturing Bacteria: Nutrition & Environment...................................................................... 35 Exercise 11 – Introduction to the Spectrophotometer ................................................................................. 39 Exercise 12 – Growth Curve of Escherichia coli........................................................................................ 43 Exercise 13 – Identification of Unknowns ................................................................................................. 51 Exercise 14 – Respiration/Fermentation/Anaerobic Respiration ................................................................ 57 Exercise 15 – Additional Tests for Gram Positive Bacteria ....................................................................... 63 Exercise 16 – Additional Tests for Gram Negative Bacteria ..................................................................... 69 Appendix A: Bacteria Used in MB 303 .................................................................................................... A-1 Appendix B: Media Used in MB 303 ....................................................................................................... B-1 Appendix C: Glossary of Media Ingredients ............................................................................................ C-1 Appendix D: Differential Stain Information ............................................................................................. D-1 Appendix E: Example Flow Chart (Graphical Dichotomous Key) .......................................................... E-1 Appendix F: Example Results Charts for Unknowns ................................................................................ F-1 Appendix G: Lab Summaries ................................................................................................................... G-1

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MB 303 Lab Schedule – Fall 2017

Week 1

Date Mon. Sept. 25, Tues. Sept. 26, or Thurs. Sept. 28

Activity

Notes Introduction to the Lab Use of the Microscope Simple Stains: Microbial Morphology Transfer of Bacteria Environmental Sampling (take-home) Mandatory 100% (online, due before next lab) Lab Notebook Due finish continue Differential Stains Pure Culture: Streak Plates introduction Scientific Writing (homework assignment) Dilution Scheme Due Assignment Due finish Quantification of Bacteria Culturing the Unculturable Lab Notebook Due Ex. 1-6, 8 (35 minutes) finish Culturing Bacteria: Nutrition & Environment Introduction to the Spectrophotometer Lab Notebook Due Finish (class discussion of results: ≈1.5 h) Identification of Unknowns (start) In-lab worksheet due before leaving lab Gram – and Gram + Flow Charts Due Growth Curve (data sheet due before leaving lab) continue Respiration/Fermentation/Anaerobic Respiration count plates (data sheet due before leaving lab) continue finish Additional Tests for Gram Positive Bacteria Additional Tests for Gram Negative Bacteria Lab Notebook Due finish

Exercise 1 Exercise 2 Exercise 3 Exercise 4 Safety Quiz Mon. Oct. 2, Week 2 Exercises 1, 2 Tues. Oct. 3, Exercise 3 or Exercise 4 Thurs. Oct. 5 Exercise 5 Exercise 6 Exercise 7 Exercise 8 Mon. Oct. 9, Week 3 Exercise 7 Tues. Oct. 10, Exercise 8 or Exercises 4 & 6 Thurs. Oct. 12 Exercise 7 Exercise 9 Mon. Oct. 18, Week 4 Exercises 3, 4, 5 Tues. Oct. 19, Midterm or Exercise 7 Thurs. Oct. 21 Exercise 10 Exercise 11 Mon. Oct. 23, Week 5 Exercises 6, 7, 9 Tues. Oct. 24, Exercise 10 or Exercise 13 Thurs. Oct. 26 Exercise 12 Mon. Oct. 30, Week 6 Exercise 13 Tues. Oct. 31, Exercise 12 or Exercise 13 Thurs. Nov. 2 Exercise 14 Mon. Nov. 6, Week 7 Exercise 12 Tues. Nov. 7, Exercise 13 or Exercise 14 Thurs. Nov. 9 Exercise 15 Exercise 16 Mon. Nov. 13, Exercises 10, 11 Week 8 Tues. Nov. 14, Exercises 12, 13, 15, Thurs. Nov. 16 16 Mon. Nov. 20/ Exercise 12 Week 9 Assignment Due Hand in as 3 separate items Tues. Nov. 21* Exercise 13 Unknown Reports due with name/seat # on each Mon. Nov. 27, Week 10 Exercises 13, 14, 15, Lab Notebook Due Tues. Nov. 28, 16 Thurs. Nov. 30 Lab Final Ex. 1-16 (1.5 h) All items are due at the start of student’s scheduled lab day, unless otherwise indicated. Items not handed in by 1 pm on scheduled lab day will be marked as late (see Course Policies on pg. vi for details about late policy). *Note: due to the Thanksgiving holiday on Thursday, Nov. 23, students in the Thursday lab section must hand in the Ex. 12 Assignment and the two Unknown Reports by Tuesday, Nov. 21, 1 pm. iii

Learning Outcomes for MB 303* After successful completion of MB 303, students will be able to: 1. Properly prepare and view microbiological specimens for examination using brightfield microscopy. 2. Use pure culture and selective techniques to enrich for and isolate microorganisms, using proper aseptic technique. 3. Estimate the number of microorganisms in a sample using both viable plate counts and spectrophotometric methods. 4. Evaluate a microbiological problem in the context of an unknown microorganism, using appropriate media-based methods for identification. 5. Accurately document and report observations and interpretations made during laboratory exercises. 6. Use appropriate microbiological lab equipment and methods, in order to conduct and analyze experimental measurements relevant to microbiology. 7. Practice safe microbiology, using appropriate protective and emergency procedures. Learner Expectations: 1. Attend lab (on time) and stay until all lab exercises are completed. 2. Read laboratory exercises in lab manual before they are to be performed. 3. Bring lab manual and lab notebook to class. 4. Come already prepared to take exams (i.e. do not wait until the night before to cram). 5. Participate in learning activities and complete tasks on time.

*All MB 303 learning outcomes have been derived from the American Society for Microbiology (ASM) Curriculum Guidelines for Undergraduate Microbiology, Part 2: Competencies and Skills, Microbiology Laboratory Skills, published Sept. 2014. http://www.asm.org/images/Education/FINAL_Curriculum_Guidelines_w_title_page.pdf

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MB 303 LABORATORY Instructors Dr. Linda Bruslind, 322 Nash, 737-1842, [email protected] (Monday and Tuesday labs) Dr. Walt Ream, 1081C ALS, 737-1791, [email protected] (Thursday lab) Office Hours During lab period or by appointment (email instructor for available days/times) Pre-requisites/Co-requisites – MB 302 Laboratory Supplies (required by each student) MB 303 Laboratory Manual (available for sale at OSU bookstore) Bound lab notebook (i.e. spiral notebook or composition book) Long-sleeved lab coat, either fabric or Tyvex (available for sale at OSU bookstore) Box of glass microscope slides (available for sale at OSU bookstore) Grading (approximate, subject to changes as necessary) EXAMS……………….….……………….…....75 pts. LAB NOTEBOOK………..……………………150 pts. UNKNOWNS………..………………………....100 pts. IN-CLASS POINTS & ASSIGNMENTS……...75 pts. TOTAL…………………………………………400 pts. Final grades are assigned on a straight percentage basis: A = 93-100%; A- = 90-92%-; B+ = 87-89%; B = 83-86%; B- = 80-82%; C+ = 77-79%; C = 73-76%; C- 70-72%; D+ = 67-69%; D = 63-66%; D- = 6062%; below 60% = F. For S/U grading a 70% (C-) or above is required to receive an “S”. Election of S/U grading should be known only to the student and their academic advisor. Format Each lab will begin with a lead-in lecture about the principles for the exercises and demonstrations of new techniques. The exercises must be read prior to class, so that students will optimize their understanding and performance during the lab period and finish during the allotted time. Some exercises require independent work while others will be done in pairs/groups. However, each student should perform their own observations, drawings, and write-ups for each exercise, to be recorded in a bound lab notebook. Ask questions when you do not understand a laboratory procedure. Good laboratory technique depends not only on knowing what you should be doing but why you should be doing it. Lab coats Each student must provide their own lab coat, to be left in the lab for the duration of the term or brought by the student each week. Students without a lab coat will be asked to leave the lab. “Loaner” lab coats are not available from the instructor. Care of Valuables Items of value should not be brought to the lab because of danger of theft or damage. The Department of Microbiology will not be responsible for personal items brought to lab. Accommodations of Disabilities Accommodations for students with disabilities are determined and approved by Disability Access Services (DAS). If you, as a student, believe you are eligible for accommodations but have not obtained approval please contact DAS immediately at 541-737-4098 or at http://ds.oregonstate.edu. DAS notifies students and faculty members of approved academic accommodations and coordinates implementation of those accommodations. While not required, students and faculty members are encouraged to discuss details of the implementation of individual accommodations. v

Course Policies  Lab safety/lab procedures: Students are expected to follow the rules for laboratory safety & procedures described in detail on pages ix-x. An online safety quiz (on Canvas) must be taken before the start of the 2nd lab. Each student may take the quiz as many times as necessary. A perfect score (100%) is mandatory or the student will not be allowed to continue in the lab.  Missed labs: Attendance is mandatory. Labs may not be made up outside of scheduled MB 303 lab times. If you have more than 1 absence, you will receive an “I” if passing the course, an “F” if not passing the course. In-class points cannot be made up.  Tardiness: Students are expected to be on time and be fully participatory for the scheduled lab time. Arriving at lab more than 10 minutes late two times will count as 1 absence. Arriving greater than 30 minutes late to lab will be considered an absence.  Attending a different MB 303 lab section: attending the other MB 303 lab section is possible only with prior permission of your instructor. Email request to your instructor as soon as you are aware of a time conflict with your scheduled lab time.  Missed exams: No make-up exams will be given. Missing an exam will constitute a zero.  Correspondence: emails sent to instructors or TAs must be done using a student’s ONID account.  Lab Notebooks: page vii describes expectations for the lab notebook.  Submission Policy: Students are allowed to submit an assignment once for grading. Components of partially completed assignments are not eligible for the late policy once the assignment has already been handed in and graded. This applies to the lab notebook and external assignments.  Late Work: all items are due at the start of a student’s scheduled lab (1 pm Monday, Tuesday or Thursday), unless otherwise indicated on the syllabus. Items handed in before the end of a student’s scheduled lab (4:50 pm) on the date due will have 10% deducted. Items handed in by 1 pm on the day following the due date will have 25% deducted. Items more than 24 hours late will not be accepted.  Grading: students have 2 weeks from the time that papers/exams are returned (or available for pickup) to contest a score. Please look your papers over carefully! Points will be updated on Canvas every 1-2 weeks by the lab TAs. Check to make sure all your grades are recorded correctly. If a grade is listed incorrectly or not posted, a student should contact their lab TA as soon as possible.  Extenuating Circumstances: exceptions to the course policies will be made only in the case of truly extenuating circumstances (i.e. serious illness, death in the family, car accident) that are documented (i.e. doctor’s note). The instructors retain the right to decide if circumstances are extenuating or not. Link to Statement of Expectations for Student Conduct: http://oregonstate.edu/admin/stucon/achon.htm Students are expected to be honest and ethical in their academic work. Academic dishonesty is defined as an act of deception in which a student seeks to claim credit for work or effort of another person, or uses unauthorized materials or fabricated information in any academic work. It includes:     

cheating- use/attempted use of unauthorized materials, information or study aids fabrication- falsification or invention of any information assisting- helping another commit an act of academic dishonesty tampering- altering or interfering with evaluation instruments and documents plagiarism- representing or presenting the words/artistry/ideas/data of another person as one's own or using one’s own previously submitted work.

When evidence of academic dishonesty comes to the instructor's attention, the instructor will document the incident, permit the accused student to provide an explanation, advise the student of possible penalties, and take action. The instructor may impose any academic penalty up to and including an "F" grade after consulting with his/her department chair and informing the student of the action taken. All persons must treat each other with dignity and respect in order for scholarship to thrive. Behaviors that are disruptive to teaching and learning will not be tolerated, and will be referred to the Student Conduct Program for disciplinary action. Behaviors that create a hostile, offensive or intimidating environment based on gender, race, ethnicity, color, religion, age, disability, marital status or sexual orientation will be referred to the Affirmative Action Office. vi

LAB NOTEBOOKS The lab notebook is an important part of your grade. Notebooks will be collected and graded throughout the term, as noted in the Laboratory Schedule. Each student must have a lab notebook designated for MB 303 alone, such as a spiral bound notebook. A lab notebook with carbon copy pages is not required. The notebook should be clearly labeled with the student’s name, lab day, and seat number on the front cover. Pages in the lab notebook should be numbered at the top. The first page of the lab notebook should be used for a Table of Contents. Each exercise should be listed, followed by the page numbers that correspond to each exercise. The table of contents must be kept up to date. Points will be deducted if your lab notebook is not neat, organized and legibly written. In a professional laboratory, a researcher's lab notebook belongs to the lab and must be written so that other people can read and follow the work. Write clearly and legibly; if your writing is illegible, you will not receive credit. Your notebook should also be well organized. If you run out of room at the end of an exercise, reference the rest of the exercise to a later page in your notebook by writing in large print at the bottom of the page, i.e. The Rest of Exercise # is on Page #. You should have plenty of room in your notebook - do not crowd things. If results will be collected later, leave several blank pages for the results. Finally, there should be no loose pages in your notebook. All inserts, such as computer graphs, should be stapled into your notebook (do not use paper clips). It is to your benefit to make sure the grader can find with ease all the pertinent information. Each lab exercise includes two parts, Objectives and Results, which should be written directly into your lab notebook. Items that need to be done outside of lab (i.e. graphs) should be well labeled and stabled into the lab notebook. Objectives should be written into your lab notebook before you start the lab. An objective is a brief statement describing the purpose of the exercise and what you will be learning about. You should know what your objectives are before you start each lab. There are examples of Objectives for Exercises 1-4 in the lab manual. Each student is responsible for writing an Objective for Exercises 5-7, 9-11, 14-16 in their lab notebook. An objective does not need to be written in the lab notebook for Ex. 8, 12, or 13. The Results Section includes all lab observations, interpretations, calculations, and graphs. All observations made during lab should be written directly into your lab notebook during lab time. Observations include not only a written description but often an interpretation or drawing of the observation as well. All drawings and descriptions should be well labeled. Therefore, a specimen viewed under the microscope should always include the total magnification, a drawing depicted as microscope field of view (i.e. a representative of what you see when looking in the microscope), and a detailed description of the relevant specimen(s) viewed. Parts of the specimen(s) should be labeled if there are parts that can be labeled, and all other pertinent information should be included. Scientific names must be written in correct binomial nomenclature (genus capitalized, species lowercase, both words underlined). Calculations and graphs can be done on your own time but must be included in the results section. You may re-write any observations that you want for your own use (i.e. re-do a table on computer outside of lab), but your grade will be based on your initial handwritten observations & interpretations made in lab and entered into your lab notebook. In order to receive full credit, be sure to use the lab time to make all the necessary observations and record all of the results that are indicated, in your lab notebook. As a final note: You should always look over your lab notebook after grading to make sure the grader did not make any mistakes and that point totals were added correctly. The policy of this lab is to give you a maximum of two weeks to get any grade changed starting the day the graded item is handed back (or available for pick up). This applies to all items in the class.

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LABORATORY SAFETY RULES & PROCEDURES 1) CLEAN desktop with DISINFECTANT (i.e. quaternary ammonium compound, QAC) at the beginning and end of class. Carefully wash hands with soap before leaving the lab. 2) KEEP YOUR WORK SPACE CLEAR – keep only the lab manual, your lab notebook, and any necessary lab supplies on your bench top; everything else should be placed under the bench, keeping the aisles clear. Backpacks/coats/cell phones etc should not be left on the lab bench top. Do not bring valuables to lab! 3) DO NOT eat, drink, chew gum or tobacco in lab. Open beverage/food containers must be left on the hallway shelf outside lab. Keep your hands out of your mouth, nose, and eyes. 4) WEAR APPROPRIATE CLOTHING – Each student is required to provide and wear a longsleeved lab coat at all times in lab. The lab coat may be left in the lab for the duration of the term or brought each week. Students without a lab coat will be asked to leave lab. “Loaner” lab coats are not available. Closed-toed shoes are required. Protective eye goggles & gloves will be provided if deemed necessary. 5) PAY ATTENTION TO ANNOUNCEMENTS - No MP3 players/iPods in lab; turn off cell phones/pagers while in lab. 6) NO UNAUTHORIZED VISITORS in the lab. NO ANIMALS in the lab. 7) KNOW THE LOCATION of the fire extinguisher (on the wall in the hallway), the fire blanket (the red box on the wall near the windows), & the eye wash (yellow capped faucet at sink). A full body shower is located outside the lab, to the right and around the corner, at the entrance to the media room in Nash 324. A partial body shower is located at the front desk in Nash 304. 8) INJURIES – report accidental cuts or burns to the instructor or TA immediately. If the injury needs professional assistance, you will be escorted to the Health Center or proper facility. 9) BROKEN GLASSWARE - Call the instructor or TA to assist you. Do not dispose of any glassware in the regular garbage cans. a. Contaminated glass is placed in the large metal can on the discard table. b. Non-contaminated glass is placed in the large cardboard “broken glass” box. 10) SPILLS – If you spill anything in lab, inform the instructor or TA so that they can assist you in proper clean-up. If culture is spilled on your clothing or belongings, they may require decontamination to assure your safety. 11) INCUBATING OF CULTURES – each workspace has an assigned number that can be used to identify your materials. Carefully label all materials to be incubated with your name/initials, seat #, and organism identification. Place materials to be incubated in the incubation tub at the front of the lab, unless otherwise directed. a. Label culture plates on agar side with your name/initials, seat #, and organism identification. Place plates in incubation tubs agar side up. This prevents moisture from forming on the inside of the lid and obliterating colonies. b. Label culture tubes on the glass (not plastic caps) with your name/initials, seat #, and organism identification. Do not use Sharpie™ to label glass items, since it does not wash off in our dishwasher. Place tubes in racks in incubation tub.

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12) BUNSEN BURNERS in the lab have almost invisible flames – use caution when they are on. Long hair must be tied back during lab to avoid contact with flame. Make sure to completely turn off your Bunsen burner when finished. Alert the instructor or TA to the smell of gas in the lab. 13) PIPETTING – Do not pipette by mouth. When using rubber pipette bulbs, insert the pipette gently into the bulb, holding the top of the pipette, to avoid breaking the pipette and potentially cutting yourself. Dispose of pipettes in the pipette collection containers. 14) DISCARDING CLASSROOM MATERIAL – All materials used in lab that are contaminated with culture (tubes, plates, pipettes, etc) must be autoclaved before cleaning or disposal. a. b. c. d. e. f. g. h. i. j.

k.

Used microscope slides should be disposed of in the cardboard box for glass waste. Used cover slips should be disposed of in the cardboard box for glass waste. Used razor blades/pins go in the metal can for contaminated glass waste. Plastic Petri plates go into an autoclave bag at the discard table. Do not discard glass items in the autoclave bag. Remove all rubber bands before discarding plates. Plastic transfer pipettes, swabs, etc go into an autoclave bag at the discard table. Do not discard glass items in the autoclave bag! Glass pipettes should be placed in the plastic containers located on the lab bench. Culture tubes go into wire baskets in metal containers (“coffins”) at the discard table. Glass bottles go into metal containers (“coffins”) at the discard table. Loosen screw caps before autoclaving. Side arm flasks should be filled with water first. Glass Petri plates go into metal containers (“coffins”) at the discard table. Uncontaminated paper towels used to clean desk with disinfectant or lens paper can be placed in metal container on the lab bench. The metal container should be emptied in the main garbage can at the end of each lab. Black wire racks are to be returned to the lab shelves. Square colored plastic racks stay on the lab bench.

15) MICROSCOPE USE – our microscopes are valuable and must be cared for properly. When replacing the microscope after use: a. Clean all oil off the 100X objective, first with flat lens paper and then flat paper saturated with lens cleaner. Wipe dry with another piece of lens paper. Residual oil will destroy the seals on the objective. Crumpled lens paper will scratch the lens. Never use anything other than lens paper to clean your microscope lenses. b. If you have gotten oil onto the 40X objective lens, inform the instructor or TA so it can be appropriately cleaned. c. Clean off dirt or oil from the stage, condensers, or oculars. d. Leave a low power (4X or 10X) objective into place. e. Wrap the cord neatly around the oculars. f. Roll out the microscope shelf completely before replacing the microscope. Sign and date the sheet in the cabinet. Have your TA check your microscope. 16) LEAVING THE LAB – Clear the lab bench of all cultures, plates, and other supplies. Empty metal can of paper waste into a large garbage can. Clean desktop with disinfectant and wash hands with soap. Make sure that you have all of your personal belongings. Please note that laboratory items (supplies, cultures, plates, VWR pens, etc) must remain in the laboratory.

Note: consistent violations of laboratory rules and procedures may result in a point penalty at the discretion of the bench TA or lab instructor.

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EXERCISE 1. USE OF THE MICROSCOPE Introduction A bright-field microscope (see Figure 1.1) is a type of light microscope that allows an observer to see a dark image against a brightly lit background using a light source in the base of the microscope. Most modern microscopes are compound microscopes, meaning that they have two types of lenses - the oculars and the objectives. These lenses provide a vital function for the microscopes - magnification or enlargement of objects. The total magnification is the product of the objective lens and of the ocular lens (i.e. multiplication of the two quantities). Magnification of each lens is indicated by markings on the lenses: typically 4X, l0X, 40X, or l00X on the objectives; l0X on the oculars. In addition to magnifying objects, microscopes enable the observer to distinguish structures that are separated by short distances. This function is called resolution. Resolution is more important than magnification. It is not always desirable to obtain the largest image possible, but it is necessary to obtain sharp detail. Resolution is dependent upon the length of the light waves used for illumination and the quality of the lenses. In bright-field microscopes, visible light with a wavelength of 400-750 nm is used. Resolution is inversely proportional to wavelength. The ordinary light microscope with oil immersion objective can distinguish or "resolve" 2 points approximately 0.2 microns apart. This is defined as the limit of resolution. To get resolution of objects smaller than this (i.e. closer than 0.2 microns) one must use an electron microscope, where a beam of electrons is used to “visualize” an object. With an electron microscope, the limit of resolution is in the range of 0.0003 microns. Another concept that is crucial to proper utilization of a microscope is its’ capacity to be parfocal. Parfocality refers to the microscope’s ability to remain in approximate focus when switching from one objective to another. Working distance refers to the distance between the specimen and objective. As you move from the 10X to the 100X objective, the working distance decreases from 8.3 mm to 0.14 mm. Of paramount importance to the microbiologist is the oil immersion objective (100X) of the microscope. Most observations of bacteria require use of the oil immersion objective. To properly use this objective, one must actually place oil on the slide surface and immerse the objective in the oil by swinging it into place. The main effect of immersion oil is to collect aberrant light rays and allow them to enter the objective. Aberrant light rays are those that would otherwise be lost due to diffraction. Without oil, light is diffracted as it passes through the glass slide into the air and not all light enters the objective. Oil has the same refractive index as glass so diffraction of the light does not occur. A phase contrast microscope is a type of compound light microscope that separates light into 2 phases that cancel each other to form an image. The phase-contrast microscope allows one to observe details in unstained living organisms. See the MB 302 text for more details on the workings of specific types of microscopes. Specimens to be viewed with a compound microscope are mounted on glass slides. A stained permanent slide is prepared by fixing whole microorganisms or sections of organisms to the slide, staining the preparation, and gluing a glass coverslip on top. Semi-permanent slides of bacteria are prepared by fixing and staining the preparation without gluing on a coverslip. An unstained wet mount is made by suspending microbes in liquid and covering the drop with a coverslip. A wet mount typically dries after 30-60 minutes, so it cannot be used over a long period of time. In addition, because of the liquid between the slide and the coverslip, a wet mount can only be observed using the 10X and 40X objectives on the microscope. However, a wet mount enables the viewer to study living microorganisms and obtain information about natural pigmentation and motility. Objectives  To become thoroughly familiar with the compound microscope and its component parts.  To demonstrate an ability to properly use the microscope using various powers of magnification.  To become adept at the preparation and observation of wet mounts.

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Figure 1.1: Typical Brightfield Microscope.

Ocular lenses (eyepieces)

Arm Objective lenses

Condenser knob

Stage

Coarse adjustment knob

Condenser Stage controls

Fine adjustment knob

Iris diaphragm Base

Light control Light source

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All microscopes in this class are compound microscopes. The basic parts and functions are: PART

FUNCTION

Ocular lenses (eyepieces)

Lenses closest to the eye, typically magnify 10X.

Ocular width adjustment

Adjust for your eye span. For the Zeiss microscopes, width adjustment is made by separating oculars to correct distance. Once this separation is made the distance between the oculars should be set on each collar to make the microscope parfocal. This adjustment must be done each time you use the microscope. The Leica and Leitz microscopes do not need this adjustment.

Revolving nosepiece

Can be rotated to change from one objective to another.

Objective lenses

Three or four magnifications: 4X (very low power); l0X (low power); 40X (high power); and l00X (oil immersion). Immersion oil used with the 100X objective has a refractive index approximately the same as glass - light passes through without being bent or lost.

Stage adjustment knobs

Moves the slide on two horizontal planes: back and forth and side to side.

Mechanical stage

Provides movable platform for slide, to change field of view.

Iris diaphragm or aperture diaphragm

Opens and closes the diaphragm to control the amount of light striking the object. Improper illumination is a common problem as the light must be increased as magnification increases.

Condenser

Condenses light waves into a pencil shaped cone, thereby preventing the escape of light waves. Its’ ideal position is just beneath the stage.

Condenser knob

Raises and lowers condenser. For this course the condenser should normally be all the way up.

Light control

Turns light source on and off. The degree it is turned on determines the intensity of light.

Light source

Uses visible light to provide illumination for sample.

Coarse adjustment knob

Moves stage up and down rapidly for purposes of approximate focusing. Should only be used when focusing with low power objectives.

Fine adjustment knob

Moves the stage up and down very minutely for purposes of definitive focusing.

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Materials per Class Hay infusion and/or pond water Microscope slides and cover slips Procedure – Work individually To remove your microscope from the cabinet roll the tray all the way out and hold the microscope in an upright position with both hands by grasping the arm and the base. Unstained Wet Mounts of Fluid Specimens 1. Prepare a wet mount of the hay infusion broth or pond water by placing a small drop of fluid on a clean microscope slide. Position a cover slip over the drop of fluid. 2. Observe, using the 4X, 10X, and 40X objectives. Wet mounts of hay infusion broth typically contain large numbers of unstained microorganisms, such as protozoa (amoebae, ciliates and flagellates), some microscopic invertebrates (rotifers and nematodes), and bacteria. Wet mounts of pond water also contain large numbers of unstained microbes, mostly unicellular or filamentous algae, some protozoa, microscopic invertebrates, and bacteria, particularly cyanobacteria. Some of the microbes move very fast. Protozoa generally are motile while algae, which may or may not be motile, are pigmented. Focusing a Wet Mount on a microscope a. Put slide on stage within movable slide holders. b. Move slide, using stage adjustment knobs, so specimen is directly over the light source. c. Move nosepiece to low power objective (l0X). d. Using coarse adjustment, move low power objective as far down as possible. e. Slowly move coarse adjustment up until you obtain best possible focus on specimen. f. Carefully change to high dry objective (40X). g. Using fine adjustment, focus until the specimen is again at maximum clarity. h. Use mechanical stage controls to move to optimal viewing areas. The fine adjustment may have to be changed as the field of view changes. i. Adjust the iris diaphragm to increase contrast. Returning microscope to the cabinet: move the nosepiece to a low power objective. Wrap the cord neatly around the oculars. Sign and date the sign-out sheet on the cabinet door. Have your TA check your microscope. Refer to "When replacing the microscope after use" of Laboratory Rules (p. viii). Lab Notebook –  Draw the observations for either a hay infusion or a pond water wet mount in your lab notebook. Draw at least 3 different organisms from the microscope field of view (i.e. what you see when you look in the microscope), observing with the 40X objective. It is not necessary to draw every cell observed – a few representative cells will suffice. Make each drawing separate, providing all the information indicated on the sample observation below. 3 pts.  Make a presumptive identification (genus level) of at least two of the organisms you observe. Examples of organisms common to hay infusion broth/pond water are depicted in Figure 1.2, Identification Guide for Pond Water/Hay Infusion Broth. 2 pts. Sample observation in lab notebook:

Type of Slide: wet mount Identification (if possible): Volvox Motility: none observed Shape: star Cell arrangement: single cells Color: bright green Total magnification: 400X

microscope “field of view”

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Figure 1.2. Identification Guide for Pond Water/Hay Infusion Broth Note: drawings are not to scale, measurements are approximate General Description Cyanobacterium, single-celled, no flagella Green algae, single-celled, no flagella

Characteristics

Picture

Very small slightly ovoid cells, green in color, nonmotile

1-1.5 μm

Small round cells, green in color, non-motile

2-10 μm

Triangular cells, green in color, non-motile

10-20 μm

Large crescent shaped cells, green in color, non-motile Green algae, multicellular or colonial growth, no flagella

Genus

Synechococcus

Chlorella

Tetraedron

Closterium 150 μm

4 or 8 cells with presence of 4 spines, green in color, nonmotile

Scendesmus 20 μm

Many egg-shaped cells contained within a mucus envelope, green in color, non-motile Fixed number of cells (4, 8, 16, or more) spherically arranged, green in color, non-motile Many small round cells contained within a mucus envelope, green in color, non-motile Disk-shaped collection of cells with hornlike projections, green in color, non-motile

Oocystis 20-40 μm Coelastrum 50 μm Sphaerocystis 20-50 μm

Pediastrum 20-50 μm

Small cells grouped in 2s and 4s in a large sac, green in color, non-motile

Tetraspora 20-50 μm

Large cells often contained in a large sac, green in color, non-motile

Chlorococcum 20-50 μm

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General Description Green algae, single-celled, with flagella

Green algae, colonial growth, with flagella

Diatoms

Characteristics

Picture

Small, single-celled, green in color (with small eye spot), 2 flagella, display rapid motility Single-celled with ridges, green in color (with eye spot), single flagella, display rapid motility

Genus

Chlamydomonas 10 μm Lepocinclis 50 μm

Cells embedded in a jelly mass with flagella pointed outward, green in color, motile

Eudorina 50-100 μm

Colony very large & hollow with cells embedded along the outer edge with flagella pointing outward, green in color, motile Boat-shaped single cells with cell face symmetry & a cross-face in valve view, golden-brown in color, slow gliding motility Boat-shaped single cells with cell face symmetry & fine lines around the valve, golden-brown in color, slow gliding motility Boat-shaped cells typically found as long ribbon-like colonies, brown in color, motile Roughly boat shaped single cells often with long narrow ends & faint lines, goldenbrown in color, motile Roughly boat shaped single cells, extremely elongated, golden-brown in color, motile The “mummy shaped” diatom, single cells with symmetry& lines, brown in color, motile

Volvox 200 μm

Stauroneis 30 x 120 μm Navicula 20 x 100 μm Fragillaria 5 x 60 μm Nitzschia 7 x 100 μm Synedra 5 x 300 μm Hemitrichia 30 x 90 μm

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General Description Filamentous algae

Characteristics

Picture

Long chains of rectangular cells with coiled chloroplasts, green in color, non-motile Long filaments composed of short cells, each containing a band-like chloroplast, green in color, non-motile Long filaments composed of brick-like cells, green in color, non-motile

Genus

Spirogyra 1 cell = 40 x 120 μm Ulothrix 10-40 μm across Microspora 1 cell = 20 x 50 μm

Long filaments usually branched composed of rectangular cells with blocklike chlorplasts, green in color, non-motile Long chains of round cells, may have heterocysts, green Filamentous cyanobacterium or blue-green in color, nonmotile Long chains of slightly rectangular cells, may have heterocysts, green in color, non-motile Long & robust filaments, individual cells difficult to distinguish, green in color, non-motile

Stigeoclonium 1 cell = 10 x 20 μm Anabaena 1 cell = 5-10 μm Nostoc 1 cell = 5-10 μm Oscillatoria 10-40 μm across

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General Description Protozoa (all singlecelled)

Characteristics

Picture

Ovoid cells covered with hair-like cilia, deep oral groove to bring in food, motile

Paramecium 50 x 200 μm

Roundish cells with short tufts of cilia, a cup-shaped mouth, with projections off rear. Motile, can walk on surfaces. Ellipsoidal cells with cilia, an oral groove and an undulating membrane, motile. Often pink tinged.

Euplotes 50 x 120 μm Blepharisma 50 x 200 μm

Ovoid cells covered with cilia, indented region leading to mouth, motile

Colpidium 30 x 70 μm

Horn shaped cells with a ring of cilia around the “bell” or mouth

Stentor up to 2000 μm

Bell shaped cells with a ring of cilia around the mouth, stalked

Microanimals (multicellular)

Genus

Vorticella 30 x 300 μm

Elongated cells with an obvious flagella extending in front Worm-like shape with segmented body, can contract, have ciliated tufts around mouth at one end Barrel shaped body with 4 pairs of stumpy legs, name means “slow walker”

Peranema 20 x 70 μm Rotifer 100-500 μm

300-500 μm

Slender worms

Tardigrade “water bear”

Nematode 100-2500 μm

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EXERCISE 2. SIMPLE STAINS: MICROBIAL MORPHOLOGY Introduction A simple stain is one in which one stain is used to create a contrast between the specimen and its background. Simple stains can be prepared quickly and give information about cell shape, size, and arrangement. The simple stain is a positive stain if the specimen is stained and not the background. To understand how a dye stains a bacterial cell, you must first know the definition of a dye. Dyes are generally salts in which one of the ions is colored. A salt is a compound composed of a positively charged ion and a negatively charged ion. The dye methylene blue is actually the salt methylene blue chloride, which dissociates as follows: Methylene blue Cl  methylene blue + + chloride

-

The color of the stain is in the positively charged methylene blue ion. Bacterial cells have a slightly negative charge when the pH of their surroundings is near neutrality. The negatively charged bacterial cell combines with the positively charged methylene blue ion, and thus the bacterial cell is stained. It is the difference in charge that produces an affinity between the dye and the bacterial cell. Dyes may be divided into two groups - basic and acidic. If the color is in the positive ion of the dye, we call it a basic or cationic dye or stain. Thus, methylene blue is a basic dye. If the color is in the negatively charged ion, we call it an acidic or anionic dye. Generally, simple stains involve the use of basic stains. Anton van Leeuwenhoek in the late 1600s first described the general morphology (shape) of bacteria as bacillus, coccus, and spiral. Apart from these differences in shape of individual cells, definite patterns in cellular arrangements are known to exist among different bacterial species, when looking at the relationship of cells as a whole. However, it should be noted that all cellular arrangements are dependent on growth conditions, and therefore, arrangement is not considered a stable characteristic. Cocci (singular, coccus), the sphere-shaped bacteria, include four important patterns for cell arrangement: pairs of cocci stuck together (diplococci), chains of four or more organisms (streptococci arrangement), irregular groups of microorganisms resembling grape clusters (staphylococci arrangement), and packets of 4 cells (tetrads). Bacilli (sing., bacillus), the rod-shaped bacteria, can be found occasionally in pairs or short chains. However, these cellular arrangements of bacilli are not as constant as in the case of the cocci, and therefore should not necessarily be considered as a characteristic for a particular species. Some bacilli do grow in long filamentous chains (filamentous bacilli) and this is an important characteristic. The spiral bacteria are found in various spiral shaped forms, as rods with one or more curves. The first group, the spirochetes, are difficult to see with a bright-field microscope since they are extremely thin (about 0.2 micrometer). They are flexible rods with several curves. The second group, the spirilla, are rigid rods possessing several curves. The third group, the vibrios, are short, curved bacteria (rods with a single curve). All three of these groups are motile. Variations in the general shapes and sizes of a bacterial species are frequently seen, and can be explained in terms of environmental factors. Pleomorphism is the term used to denote these variations within a species. Objectives  Observe simple stains of bacterial smears  Identify basic bacterial shapes and cell arrangements  Differentiate size differences among different microbes

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Materials per Student Prepared slide: Spirillium, Bacillus subtilis, or Staphylococcus aureus Procedure Stained Permanent Mounts of Organisms 1. Each student needs to get 1 prepared slide in focus under the 100X objective lens. Show your TA the organism that you got in focus. 2.5 in-class pts. 2. Observe each of the three prepared slides of different bacteria (Spirillium, B. subtilis, and S. aureus) by working with other students near-by. 3. At the end of lab the oil should be cleaned off each slide using a paper towel. Focusing a Stained Slide on the microscope a. Put slide on stage within movable slide holders. b. Using mechanical stage controls move slide so that a promising area is directly over the light source. c. Move nosepiece to low power objective (l0X). d. Using coarse adjustment, move low power objective as far down as possible. e. Slowly move coarse adjustment up until best possible focus on specimen is obtained. f. Move the 40X objective into place and focus with fine adjustment. If you lose focus at this point, make sure your slide is right side up and your condenser is directly beneath the stage. g. Swing the 40X objective out of the way, but do not snap the 100X objective into place yet (and do not make any other adjustments on the microscope). Squeeze a small drop of oil onto the slide, directly over the path of light on the specimen. h. Carefully switch to the oil immersion lens (l00X). If your microscope is parfocal, the oil immersion lens should just barely touch the oil on the specimen. Never go back to the 40X objective once oil has been added to the slide. If this occurs, let the TA know so your 40X objective can be removed for special cleaning. i. Use only the fine adjustment knob to adjust focus for maximum clarity. Very little focusing should be needed at this point. If you do not get focus easily, check with a TA. j. Use mechanical stage controls to move to optimal viewing areas. The fine adjustment may have to be changed as the field of view changes. k. Always remember to clean the oil immersion lens, and the condenser, if needed, with lens cleaner and lens paper when one is finished. A MICROSCOPE SHOULD NEVER BE PUT AWAY WITH OIL ON IT. Lab Notebook.  Draw your observations for all 3 organisms in your lab notebook. Note: All drawings in your lab notebook should be completely labeled (see example below). Be sure to label the full scientific name of each organism, using correct binomial nomenclature: Capital genus, lowercase species, both underlined. Indicate the shape, color, cell arrangement, and approximate size of each organism (for purposes of estimating size, assume Staphylococcus aureus is approximately 1 m in diameter). Record the total magnification for each observation (remember that total magnification is the magnification of the objective lens multiplied by the magnification of the ocular lens). 3 pts. Sample observation in lab notebook: (showing microscope field of view)

Type of Slide: simple stain Organism: Vibrio cholerae Shape: spiral (curved rods) Cell arrangement: single cells Approx. size: 1 x 4 µm Color or stain used: methylene blue Total magnification: 1000X

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EXERCISE 3. TRANSFER OF BACTERIA Introduction Organisms are present all around us and when microbes are transferred from one medium to another it is important to exclude these environmental organisms. The introduction of microorganisms into a culture medium is termed inoculation. In order to cultivate only those microorganisms that are desired, aseptic technique must be employed while transferring organisms from a viable culture to a sterile medium. This technique will be demonstrated for this exercise. Aseptic technique is the transfer of culture and media without contamination by unwanted organisms. Aseptic technique is just a matter of common sense. Unless a surface is specifically sterilized, it is contaminated with microorganisms (disinfectants do not sterilize). If a sterile surface comes into contact with a non-sterile object that surface is no longer sterile. A few aseptic techniques: 1. Flame inoculating loops and needles to a cherry red to sterilize and prevent environmental contamination. Flame the entire part of the loop/needle that will contact culture and/or medium. 2. Flame mouth of all glass tubes and bottles immediately after lid is removed and before the lid is replaced. This sterilizes the mouth of the tube/bottle and creates an updraft of warm air that will move contaminants away from the open container. 3. Never turn a lid upside down or set on the table top since this will allow debris and microorganisms to fall into the lid and consequently into the culture. Lids should be held in your hand with the inside facing down until it is replaced. 4. Hold all tubes at an angle to prevent debris and microorganisms from falling into the tubes. 5. Inoculating loops, inoculating needles, swabs, pipettes, and syringes that are employed in the mechanical transfer of the cultures should be sterile to prevent environmental contamination. 6. Sterile pipettes should not be touched at the tip or come in contact with any surface, to prevent environmental contamination. Solid and broth media are employed to grow microbes in the laboratory. Broth medium becomes turbid (cloudy) when microbes grow in them. Broth cultures are easy to work with and allow all the microbes access to nutrients, but it’s impossible to tell if the culture is pure or not. In addition, oxygen does not easily diffuse into a broth medium, although oxygen can be introduced by putting the cultures in a shaking incubator. Agar slants are test tubes containing solid culture media that were left at an angle while the agar solidified. Agar slants provide a large aerated surface for microbes to grow on and do not dry out quickly, allowing cultures to be kept for extended periods of time. Contamination may or may not be visualized depending upon how different the contaminant is from the organism of interest. Agar deeps are formed by allowing agar to solidify in the bottoms of test tubes. Oxygen does not easily diffuse into these agar deeps, providing an ideal environment for anaerobic microbes. However, it can be difficult to extract microbes from an agar deep, in order to do additional testing. Agar plates provide a large surface area to grow and visualize microbes, and can be used for a variety of samples. Unfortunately, stored plates dry out very quickly or become contaminated over time. Plates should always be incubated “agarside up,” to prevent condensation from dripping back down onto microbial growth on the agar surface. All items for incubation should be properly labeled, to provide basic contact information and organism identification. In MB 303 that means student name/initials, seat #, and organism name. Temperature should also be indicated, if something other than the standard 37oC will be used. Plate dilution should be indicated if the agar plate is part of a dilution series. Agar plates should always be labeled on the base (as opposed to the lid) to insure that information remains associated with the microbial growth. Tubes should be labeled on the side of the tube (as opposed to the lid), for the same reason. Objectives  Recognize the reasons for aseptic technique  Practice aseptic transfer methods  Employ different forms of microbiological media

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Materials per Pair 1 Plate Count Agar (PCA) plate 4 tubes containing brain heart infusion (BHI) broth 2 tubes BHI agar slant 2 tubes semisolid BHI agar deep Broth cultures: Micrococcus luteus, Escherichia coli Procedure (see Figure 3.1) – Work in pairs 1. Remove the cover of a PCA plate and allow it to sit open until the end of lab. Alternatively, you and your lab partner can gently touch the agar surface, transferring some of the microbes from your fingers to the agar. Label the plate appropriately with name/initials, seat #s, and sample obtained (i.e. “Air Microbes” or “Finger Microbes.”) Be sure to label on the base and incubate the plate “agar-side up.” 2. Set up an aseptic technique control. This will check your aseptic technique to demonstrate that you can transfer a loopful of sterile medium from one tube to another without contaminating it. Each member of a pair should transfer a loop of sterile uninoculated BHI broth into another tube of sterile uninoculated nutrient broth using the following procedure: To transfer BHI broth, hold the inoculating loop in your dominant hand and hold one of the broth tubes in the other hand. a. Sterilize the loop by holding the wire in a Bunsen burner flame. Heat to redness. b. Holding the loop like a pencil, curl the little finger of the same hand around the cap of the broth tube. Gently pull the cap off the tube. Do not set the cap down, as the lab bench is not sterile. c. Holding the tube at an angle, pass the mouth of the tube through the flame. Always hold culture tubes and uninoculated tubes at an angle to minimize the amount of dust that could fall into them. d. Immerse the sterilized, cooled loop into the broth to obtain a loopful of medium. You may cool the loop by touching it to the inside of the tube above the medium. Flame mouth of tube and recap. e. Remove the cap from the other tube of sterile BHI broth as previously described and flame the mouth of the tube. Immerse the inoculating loop into the sterile broth. Flame the mouth of the tube and replace the cap. Return the tube to the test tube rack. f. Re-flame the loop until it is red and let it cool. Some individuals prefer to hold several tubes in their hands at once. Do NOT attempt holding and transferring between multiple tubes until you have mastered aseptic transfer techniques. g. Place the "inoculated" control tube (appropriately labeled) in the incubation tube at the front of your lab bench. Label directly on the glass; do not mark on tube caps! 3.

Following the above procedure (Steps 2a-2g), each member of a pair should inoculate one of the two provided organisms. One partner inoculates E. coli, the other M. luteus, into a tube of sterile BHI broth. This is the same BHI broth from which you removed a loopful of medium in step 2.

4.

Using the same organism as in #3, each member of a pair should inoculate a BHI agar slant. Repeat steps 2a through 2d and inoculate the slant by moving the loop gently across the entire agar surface from the bottom of the slant to the top, being careful not to gouge the agar. Flame the mouth of the tube and replace the cap. Flame your loop and let it cool.

5.

Using the same organism as in #3, each member of a pair should inoculate a BHI agar deep. Dip an inoculating needle into the broth culture and then into the agar deep by plunging the needle straight down the middle of the deep to the bottom of the tube, then pull out through the same stab. Use the same aseptic technique as described in 2a to 2c. Flame the mouth of the tube and replace the cap. Flame your needle and let it cool.

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6.

Place all inoculated tubes plus the control into the 37° incubation tub for your lab bench. Make sure that all tubes are fully labeled with initials, seat number, and organism.

7.

After incubation, observe and record the growth on the agar surface of the PCA plate, noting differences in colony appearance, such as texture, color, and size. Note whether your control tube is still sterile (indicated by a lack of turbidity). Observe and record the growth in/on the BHI slant, deep, and broth, for both E. coli and M. luteus.

Figure 3.1: Culture Transfer Using Aseptic Technique.

Lab Notebook (observations after growth, Week 2)  Record the number of colonies that grew on your PCA plate. If one had no colonies on their PCA plate, could they assume the environment is sterile? Explain why or why not. 1 pt.  Record the results for the control tube that you set up. If you observe any turbidity indicate what you might do differently next time to improve your aseptic technique. 1 pt.  Record the results of inoculated tubes (broth, slant, deep) in your lab notebook, in terms of amount of growth, location of growth in tube, and pigment production, for both E. coli and M. luteus. 3 pts.

13

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EXERCISE 4. ENVIRONMENTAL SAMPLING Introduction The microbial contamination of objects and surfaces is of considerable interest to individuals in a number of occupations. Such occupations include people who are health inspectors, food processors, hospital epidemiologists, and clean room technicians, just to name a few. These people need to be able to judge accurately the degree of cleanliness of many items and areas, thus ascertaining what hazards of infection and/or contamination, if any, remain. In order to accomplish this, several techniques have been devised. For the sampling of surfaces, RODAC plates are commonly used to obtain a direct environmental sample, where RODAC stands for Replicate Organism Detection and Counting. The RODAC plate is a special plate in which the agar has been slightly overfilled due to a higher concentration of agar in the medium preparation. When the cover is removed, it can be seen that the agar surface protrudes slightly above the lip of the plate, providing a convex surface. In use, this agar surface is pressed lightly on the surface being sampled, the lid is replaced, and the plate is incubated. Organisms on the surface sampled adhere to the agar and grow as discrete colonies which may then be counted. Thus, an approximate idea as to the degree of sanitation can be obtained. One limitation of the RODAC plate is that it can only be used successfully on flat, impervious surfaces which are free of crevices. In addition, research has shown that the RODAC plate only removes ≈0.1% of contact flora, much less than the swab technique described below. A second method that has the distinct advantage of being useful on irregular surfaces is the moist swab technique or swab contact method. Here a sterile swab is moistened with a special phosphate buffer diluent solution and is then used to wipe a defined area on objects or surfaces. The swab is then returned to the vial containing the diluting fluid, tightly capped, and transported back to the laboratory. The diluent keeps the organisms in an osmotically correct environment so that they remain viable during transport back to the laboratory; however, nutrients are not available so growth does not occur. Once at the lab, an aliquot of the fluid is plated out on appropriate medium, providing a solid surface on which to observe the microbial growth following incubation. Individual colonies are observed and counted. The colony count is adjusted to represent the number of microbes per milliliter, and then used to determine if the surface or surfaces that were sampled were adequately cleaned. It is recommended that each laboratory establish its own values for what constitutes an adequately cleaned object or surface, based on guidelines for their field. For example, the parameters for a surgery suite might be more rigorous than the parameters for a restaurant bathroom. In this class we will use the standards recommended by the U.S. Public Health Service when performing environmental sampling of a commercial food preparation area. For the RODAC plate acceptable sanitation is defined by samples that have not more than 100 colonies. Samples that have 100 or more colonies are deemed to have unacceptable sanitation. The same parameters are used for the swab technique, after adjustments have been made for volume plated (microbial counts are adjusted to reflect the number of microbes in a milliliter). It is important to note that the quantity of microbes associated with a surface is not always an accurate parameter about the possible danger associated with those microbes. The microbial identity is also of importance and may be more significant than numbers alone. Objectives  Employ 2 different surface contact methods for environmental sampling.  Evaluate whether a surface meets the recommended sanitation levels prescribed by the U.S. Public Health Service  Recognize how to compare the number and types of microbial colonies with the degree of sanitation.

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Materials per Student RODAC plate 1 Hardy Diagnostics™ neutral buffer with swab 1 PCA pour 1 sterile Petri dish Hockey stick + 95% ethanol Sterile and unsterile 1 ml pipettes Procedure - This exercise takes place over three laboratory sessions – Work individually First Lab Period: collect a paper bag with the needed supplies in it. To Do at Home Before the Second Lab Period: A. RODAC plates 1. These plates are to be used to sample a flat location, such as a cutting board or countertop. 2. To inoculate the RODAC plate, remove lid, press agar firmly, but gently, onto surface being sampled using a slightly rocking motion. Replace lid and tape to prevent lid from falling off. Label the plate on the bottom as to site sampled and your name and seat number. Bring your RODAC plate back to class on the date specified by instructor. You should inoculate these plates as close to lab time as possible. B. Swab + Buffer 1. Select 4 similar surfaces to swab. The swab technique is typically used to sample 4 similar surfaces, such as 4 spoons or 4 glasses, in 2.5 cm2 swatches. 2. Unscrew cap and press excess buffer from the swab on the side of the tube. Rub swab back and forth on an area ≈2.5 cm2. 3. When finished swabbing the first surface, immerse the swab back into the buffer, swirl, and press out against the side as before. Using the same swab, repeat on another object or surface until a series of 4 similar surfaces have been swabbed. 4. After the fourth surface has been swabbed, put the swab back into the vial of buffer and tightly cap. Label with what type of surface was swabbed. During the Second Lab Period: A. Label RODAC plate with name/initials, seat number, and surface sampled. Place the RODAC plate in the appropriate incubator box agar side up to prevent condensation that forms on the inside of the lid from dripping onto the agar and disrupting the colony formation. RODAC plates will be incubated at 37°C for 48 h. B. Swab + Buffer 1. Obtain a melted, tempered tube of PCA agar from the 50°C water bath (agar melts at 100oC but does not re-solidify until 45oC). Wipe the outside of the tube with a Kimwipe. Remove cap, flame the lip of the tube and pour contents into a sterile Petri dish. (When pouring agar into the Petri dish, lift up the top of the Petri dish on one side. Do not completely remove the top.) Let solidify. 2. After the agar in the Petri dish has solidified, remove 0.2 ml aliquot of the buffer in the vial using a sterile pipette and pipette bulb (only 0.1 ml will be used to prepare the plate, but it’s best not to use the last 0.1 ml in a pipette). Practice with water and a non-sterile pipette first. Directions for using a pipette bulb: a. See Figure 4.1. Insert pipette gently into bulb. Squeeze the top button “A” with your thumb and index finger while you compress the bulb with other three fingers and palm. This creates a vacuum that can be used to suck liquids into the attached pipette. If you squeeze the bulb without pressing this button, valves in the pipette bulb are displaced. b. Put pipette into liquid you wish to dispense. Gently press button “S” to draw liquid into pipette. Do not apply excessive pressure as the fluid may flow into the pipette bulb, contaminating it. (If liquid does get in the bulb notify your TA or instructor so that the bulb may be disinfected.)

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Figure 4.1. Pipette bulb. c. To release liquid from pipette, press button “E”. To completely empty pipette, place your finger over the hole on the E arm and press toward the tiny bulb on the E joint. This will expel out the last bit of liquid in the pipette 3. Expel 0.1 ml onto the surface of your solidified agar plate. The remaining 0.1 ml can be expelled into the pipette discard container. 4. Sterilize an aluminum "hockey-stick" by dipping the stick into a bottle of 95% ethanol. The stick is then touched to a Bunsen burner flame to ignite the ethanol. Do not hold the stick in the flame. Cool stick by touching to the agar where fluid was not dispensed. Use the stick to evenly spread the fluid across the agar surface. After spreading, the stick should be re-sterilized by dipping in ethanol and touching to flame to ignite the ethanol. 5. Label PCA plate with name/initials, seat number, and surface sampled. Place plate in appropriate incubation box agar side up. PCA plates will be incubated at 37oC for 48 h. Third Lab Period: A. RODAC plates: Count the colonies on your plates. A plate with less than 100 colonies indicates a surface with acceptable sanitation. A plate with 100 colonies or more indicates a surface with unacceptable sanitation. If a colony count cannot be obtained because there is solid growth across the agar surface or there are too many colonies to distinguish individual colonies, sanitation is definitely not acceptable. Record the colony count in your lab notebook. Indicate if sanitation was acceptable or not. B. Swab + Buffer: Observe PCA plate for colonies following incubation. Multiply the colony count by 10 since only 0.1 ml was plated and acceptable sanitation levels are standardized to 1 ml for this method. Acceptable plates are those with less than 100 colonies after standardization. Record the colony count in your lab notebook. Indicate if sanitation was acceptable or not. 1. After counting colonies on each plate, select a colony from either plate to Gram stain. Be sure to add a small drop of water to your clean slide before adding a small amount of material obtained from an isolated colony. Thoroughly mix the microbes in with the water before spreading over the entire surface of your slide, creating a thin smear. Let air dry, heat fix, and then proceed with the steps of the Gram stain. 2. Observe and record observations in your lab notebook. 3. Show your TA your Gram stain, in focus under 100X. 2.5 in-class pts.

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Lab Notebook  Record your colony count results for both methods. 2 pts.  Indicate if sanitation was acceptable for each surface/surfaces sampled. 2 pts.  Record observations for Gram stain. In addition to the typical microscopic observations listed in Exercise 2 (see “sample observation in lab notebook”), you should also indicate the Gram reaction (positive or negative). 1 pt.  Record colony appearance for the colony that was Gram stained, describing the five colony characteristics listed in Table 6.1. 1 pt.  Use Table 4.1 and your observations to determine the presumptive identification of the colony that was Gram stained. 1 pt.  Research the organism (or organismal group) identified from your colony. Explain whether or not this organism would cause you any concern about the surface(s) that you sampled. Why or why not? Provide complete reference information, using ASM style (described in Ex. 8). 2 pts.

Table 4.1. Physical Characteristics of Common Environmental Microflora Organism/ Group

Gram reaction/morphology

Colony description

Bacillus sp.

Gram + bacilli (large) in chains

Large, white, textured

Coliform

Gram – bacilli (small)

Medium, tan or translucent, smooth

Lactobacillus sp.

Gram + bacilli (small-med)

Medium, white or tan, smooth

Micrococcus sp.

Gram + cocci/tetrads (small)

Small, yellow/white/pink, smooth

Neisseria sp.

Gram - diplococci

Small, grey or yellowish, smooth

Pseudomonas sp.

Gram – bacilli (small-med)

Medium, tan/cream/white, viscous

Staphylococcus sp.

Gram + cocci (medium)

Medium, white, smooth

Medium, pink/yellow/orange/cream, pasty in appearance *Yeast, as fungi, have a cell wall made of chitin and thus are not considered eligible for classic characterization using the Gram stain. However, they do tend to retain the initial crystal violet stain, resulting in a purple or “gram positive” appearance. Yeast

“Gram +” ovals (large)*

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EXERCISE 5. DIFFERENTIAL STAINS Introduction A simple stain can be used to determine morphology, cell arrangement, and cell size. If we wish to learn more about the cell, such as whether it contains an endospore or possesses a capsule, we must resort to a differential stain. A differential stain typically employs the use of two (or more) dyes to distinguish organisms based on cell characteristics or structures. The Gram stain is a differential stain commonly used in microbiology. It is one of the first steps in the differentiation and identification of a bacterial species; it distinguishes between the two main cell wall types found in bacteria. The Gram stain technique was developed in 1884 by Christian Gram. The Gram stain requires the use of four reagents. The first dye added to the smear is crystal violet, which is followed by an iodine solution. The iodine acts as a mordant (a specialized term used in dyeing), combining with the crystal violet to form an insoluble colored compound in the bacteria being stained. This insoluble precipitate is called the crystal violet-iodine complex. After decolorizing with acetone-alcohol a safranin counterstain is applied. Bacteria that resist decolorizing and retain the crystal violet-iodine complex appear purple and are called gram positive. Conversely, those cells that decolorize or lose the crystal violet-iodine complex more rapidly will accept the safranin counterstain and appear pink/red. These are the gram negative bacteria. The Gram stain results from structural differences in the cell walls of gram positive and negative bacteria. Gram positive bacteria have a thick homogeneous layer of peptidoglycan while gram negative bacteria have a thin peptidoglycan layer plus an additional outer membrane. The peptidoglycan acts as a barrier, preventing the loss of crystal violet, the first dye applied in the gram stain. The second step is the application of Gram’s iodine, which acts as a mordant, chemically combining with the crystal violet to form a larger complex. This encourages retention of the crystal violet. The third step, the decolorizing step, is the application of acetone-alcohol. Acetone-alcohol causes the thick peptidoglycan layer of the gram positive bacteria to shrink, locking in the crystal violet. The alcohol dissolves some of the lipids from the lipopolysaccharide (LPS) layer of the gram negative bacteria, leading to larger pores and loss of the crystal violet-iodine complex. The last step, application of the counterstain safranin, stains the now clear gram negative bacteria a pink color so that they can be visualized. The differentiation of the gram reaction is not an absolute, all-or-none phenomenon. A number of factors can result in inaccurate gram reactions, such as the following: 1. Improper heat fixing of the smear. If a smear is heated too much, the cells can rupture. If the cells are ruptured, the gram reaction will be undeterminable for either gram+ or gram- bacteria. 2. Cell density of smear. An extremely thick smear may not colorize/decolorize as rapidly as one of ordinary density, causing mixed results (cells staining both pink and purple) or false reactions. 3. Length and thoroughness of washing after crystal violet. If CV remains on the slide after I2 is added, a false gram+ result can be obtained because CV will combine with I2 and precipitate. 4. Age of bacterial culture. Gram reactions are typically reliable for cultures 24 hours old or less. Gram+ cultures older than 24 hours have increased breaks in their cell wall and thus may convert from an original gram positive reaction to a gram negative reaction. Often a mixture of gram+ and gram– cells appear in the microscope. This is called a gram variable reaction. There are many other types of differential stains, in addition to the Gram stain. An acid-fast stain is used for bacteria with a high mycolic acid content in their cell wall, making them resistant to the decolorizing step of the Gram stain. A capsule stain highlights the polysaccharide layer that some bacteria have, using an acidic stain on the background and a basic stain on the cell, leaving the capsule as an unstained halo around the cell. An endospore stain uses a heating step to drive one stain into the endospore coat to allow identification of the bacterial endospores, while using a second stain on the vegetative cells. See Appendix D for more details about the additional differential stains to be observed in lab. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms Gram stain reagents (crystal violet - Gram's iodine - acetone-alcohol - safranin) Broth culture: mixture of Escherichia coli (gram neg) & Staphylococcus aureus (gram pos) Demo slides of additional differential stains: acid-fast, capsule, endospore Procedure – Work individually A. Slide preparation: 1. Clean your slide. Before one can stain bacteria, a bacterial "smear" must be made. In order to do this successfully, it is imperative to begin with a clean glass slide. If the slide is not thoroughly cleaned, the bacteria will not adhere to the surface of the slide and will wash off during the staining procedure. To properly clean a slide, thoroughly wash with dish detergent, rinse with water, blot dry with a paper towel and lightly flame with the Bunsen burner. This last step is essential to insure that all traces of grease or oil have been removed from the slide, since bacteria will not adhere in such areas. After flaming, avoid touching the slide surface or oils will be redeposited on the slide. Hold the clean slide by the edges. 2. Make a smear. Prepare a bacterial smear from the broth culture by thinly spreading a loopful of culture over the entire surface of the clean slide (a smear is prepared from a solid culture by mixing a small amount of a bacterial colony with a small amount of water on the slide surface). See Figure 5.1. Remember to sterilize the loop before picking up bacteria and always keep your smear bacteria side up during the heat fixing and staining process. 3. Heat fixing the smear. First, allow the bacterial smear to air dry. If the bacterial smear is not completely dry when passed through the flame, the liquid will boil and the bacteria will explode. Next, pass the slide, bacteria side up, through the Bunsen burner flame once or twice at a medium speed. If a smear is overheated, the integrity of the cells will be destroyed and the bacteria will appear shapeless and enlarged. The slide should never get too hot to hold. This last step serves two functions: it kills the bacteria, and it denatures the proteins in the cells thereby fastening or "fixing" the organisms to the slide. B.

Gram stain procedure: 1. Each student should prepare a bacterial smear of the mixed culture containing both E. coli and S. aureus, using the directions above. Air-dry and heat-fix the smear. 2. Gram stain the bacterial smear in the following manner: a. Cover bacterial smear with crystal violet for 1 minute. Thoroughly rinse with water, shaking off the excess at the end. Do not dry. b. Cover bacterial smear with Gram’s iodine for 1 minute. Thoroughly rinse with water, shaking off the excess at the end. Do not dry. c. Decolorize the smear by holding the slide at an angle over the sink and applying acetonealcohol drop by drop until the drops coming off the bottom of the slide are very light purple in color. If the smear was made from a broth culture it will typically take 3-6 drops of acetone-alcohol. If the smear was made from solid culture (i.e. a colony) it will take more acetone-alcohol, due to the thickness of the smear. Thoroughly rinse with water, shaking off the excess at the end. d. Counterstain smear with safranin for 1 minute. Thoroughly rinse with water. e. Blot dry with a paper towel. Do not rub. 3. Observe slide with oil immersion lens (100X) of the microscope, after first focusing with the 10X and 40X lenses. Refer to Ex. 2 for focusing directions Record observations for both bacteria, including Gram reactions, morphology, approximate size, and cell arrangement. If the Gram stain did not turn out, refer to Table 5.1 for a trouble-shooting guide. 4. Show your TA the slide in focus under 100X and identify the Gram reaction and morphology of each organism. 2.5 in-class points.

C.

Additional differential slides 1. Observe the 3 differential slides set up as demos: acid-fast, capsule, endospore. Record observations in your lab notebook, as microscope field of view. Be sure to label each drawing with as much detail as possible.

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Figure 5.1. Preparing Bacterial Smears.

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Lab Notebook  Record observations for your Gram stain. In addition to the typical microscopic observations listed in Exercise 2 (see “sample observation in lab notebook”), you should also indicate the Gram reaction (positive or negative). 1 pt.  Record observations of the additional prepared differential stains (acid-fast stain, bacterial capsule, and endospores), as microscope field of view. Label with as much information as possible. 3 pts. Note: The basis and procedure for each of these additional stains is provided in Appendix D.

Table 5.1. Trouble-Shooting Guide for Gram Stains Observation under Likely explanation(s) the microscope Cells lack definite shape or Cells ruptured - bacterial smear morphology not fully dried when heat fixed Cells ruptured - bacterial smear over heated when heat fixed No obvious cells on slide

Cells washed off during staining protocol Slide is upside-down

Large dark purple/black crystallike objects are present Small blue or purple dots are present (slightly different plane of focus than bacteria) Gram positive bacteria appear pink Gram negative bacteria appear purple

Crystal violet artifact - slide was hot when CV was applied Crystal violet precipitate – CV was not fully rinsed off before iodine was applied Smear was over decolorized

Both pink and purple cells are present (i.e. gram variable reaction)

Smear was under decolorized Crystal violet precipitate – CV was not fully rinsed off before iodine was applied Culture is contaminated

Some areas of the smear were over or under decolorized Bacterial culture is old Cells are blue in color

Methylene blue used instead of crystal violet

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Solution(s) Allow bacterial smear to completely air-dry before the heat fixing step Pass the bacterial smear only 1-2 times through the flame, at a rapid rate Heat fix smear before staining slide Always keep slide bacteria side up during staining process Do not over heat fix – pass slide 1-2x through flame at rapid rate Fully rinse off CV before applying iodine Decolorize carefully; always make a thin even bacterial smear Decolorize carefully; always make a thin even bacterial smear Fully rinse off CV before applying iodine Be sure to use good aseptic technique; if source is a plate then select isolated colonies for smear Make a thin even bacterial smear Use the freshest possible culture for Gram stain Look for the “green metallic sheen” indicative of CV on the dye bottle

EXERCISE 6. PURE CULTURES: PREPARING STREAK PLATES Introduction A pure culture is a group of bacteria that is of one species. Pure culture techniques for bacteria were developed in the early 1880's by Robert Koch, a German microbiologist. Meat extracts were used to provide nutrients for the bacteria. At the suggestion of Fannie Hesse, the wife of one of Koch's assistants, agar was implemented as a solidifying agent for broth media. This was a large improvement over the gelatin that had been used as a solidifying agent as agar is not easily degraded and does not melt at low temperatures. Richard Petri, another of Koch's assistants, developed the Petri dish, a shallow lidded container used to hold the agar medium. The techniques presently used for the isolation of bacteria and the maintenance of pure cultures were thus established over 100 years ago. Medium used to culture microorganisms is sterilized in order to prevent the growth of contaminants introduced during the process of its preparation. The most common way to sterilize medium is by autoclaving (steam under pressure) at a temperature of 121oC at 15 lbs of pressure/sq. in. for a period of 15 minutes. One of the most common methods employed in the laboratory to obtain a pure culture is the preparation of a streak plate. To properly streak a plate for isolation means spreading out the organism(s) by means of the inoculating loop, or thinning the culture, until single colonies result. Theoretically, each single colony consists of a cluster of cells that originate by cell division from one bacterial cell. Thus, each colony, if well isolated from other colonies, does represent a pure culture of bacteria. Streak plates are used to isolate a pure culture when more than one culture is present and also to verify that a particular culture is pure. This lab will utilize a three-phase streaking pattern, also known as the T-streak. The plate is divided into three equal sections, marking with a pen on the bottom of the plate. A loopful of bacteria is aseptically picked up with a sterile inoculation loop and then deposited in the top section of the plate by dragging the loop back and forth on the agar surface. This is known as the area of heavy inoculum, since bacterial growth should completely cover this area after incubation. The loop is then re-sterilized, cooled, and used to drag a small amount of bacteria from the area of heavy inoculum into the 2nd section of the plate, followed by a discrete zig-zag pattern to thin out the bacteria on the agar surface. This section should have much less growth than the first section. The loop is re-sterilized again, cooled, and used to drag a small amount of bacteria from the 2nd section into the 3rd section of the plate, once again followed by a discrete zig-zag pattern. The last section should have the least amount of growth, thus yielding isolated colonies. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms per Student Broth culture: mixture of Escherichia coli and Staphylococcus aureus 1 trypticase soy agar (TSA) plate Procedure – Work individually 1. After observing the correct procedure for streaking a plate and referring to Figure 6.1, each student should prepare a streak plate of a mixed culture containing E. coli and S. aureus, using good aseptic technique. Label plate with your name/initials, seat number, and “mixed culture.” 2. Invert and place plate in the appropriate incubation tub, to be incubated at 37°C for 48 hr. 3. After incubation, observe the growth on the plate that you prepared. Note whether you obtained isolation of both bacteria or not, as well as specific growth characteristics of isolated colonies (See Table 6.1 on next page).

Figure 6.1: Streak Plate: Three-Phase StreakingPattern.

1. INITIAL STREAK Pick up bacteria using a sterilized inoculation loop and streak them thickly on the top ⅓ of your plate.

1

3

2

Rotating your plate 90o between streaking each section will keep the streaking pattern in the correct orientation.

2. SECOND STREAK Flame the loop. Touch the hot loop to an unused portion of the agar to cool before going across the previous streak once, then streaking section 2 in a nonoverlapping zig-zig fashion.

3. THIRD STREAK Flame the loop. Cool and streak from section 2 to 3, only going once across your second streak. Streak section 3 in a non-overlapping zig-zag fashion. Be sure to fully sterilize your loop between each section, in order to thin the bacteria.

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Lab Notebook  Observe and record the results of your plate. Describe what each colony type looks like, using Table 6.1 to indicate individual characteristics for each category (size, pigmentation, form, margin, and elevation). 2 pts.  Indicate if you obtained isolation for both of the bacteria, only 1 bacterium, or neither. Provide several possible reasons why one might not obtain isolation (for one or both organisms). 2 pts. Table 6.1: Characteristics used for Colony Evaluation on Agar Plate Size

pinpoint, small, moderate, large

Pigmentation

color of colony – can include white, cream, tan, yellow, pink, red, etc. A non-pigmented colony would be transparent.

Form (shape of the colony) Circular

Irregular

Filamentous

Rhizoid

Margin (appearance of colony outer edge) Entire

Lobate

Undulate

Serrate

Filamentous

Elevation (degree to which colony growth is raised on agar surface) Flat

Raised

25

Convex

Umbonate

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EXERCISE 7. QUANTIFICATION OF BACTERIA Introduction Bacteria are extremely small and often reproduce very quickly. It is not unusual for them to grow to a concentration of 1 billion (109) bacteria/ml. In order to obtain an accurate count of the number of living bacteria in a liquid culture, the culture must typically be diluted first. One method of determining the number of bacteria in a suspension is to perform a viable plate count, where dilutions of a culture of bacterial cells are prepared. Aliquots of the dilutions are plated onto plate count agar with sterile pipettes and spread with hockey sticks, incubated at 37°C, and number of colonies counted. Not all bacterial cells produce colonies, however, as some bacteria tend to clump or aggregate. For this reason, results are reported as colony forming units (CFU)/ml of a bacterial culture. Ideally, only plates with 25-250 colonies are used in the determination of a bacterial concentration. Counts above 250 are considered to be Too Numerous To Count (TNTC) because it is impossible to tell if colonies are truly separate. Plates with less than 25 colonies have been shown experimentally to have very poor accuracy in predicting bacterial concentration. A single contaminant could greatly skew the final concentration value. When the number of bacteria is unknown, a wide range of dilutions is prepared and plated. In this way you will have at least 1 or 2 plates within the countable range (25-250) to count and use in your calculations. If more than one plate is in the countable range the CFU/ml for the original bacterial suspension is determined by averaging values together to produce a single CFU/ml. It is important to remember that the various dilutions prepared for a viable plate count all derived from a single, original suspension of bacteria. The goal is to combine the most accurate data to determine the concentration of the original bacterial suspension. In order to obtain accurate data, it is imperative that good aseptic technique be used. Remember, the pipettes that you will be using are sterile - until you open the can. It is acceptable to leave the can open while you are working but it should be closed as soon as you finish a dilution scheme. Do not lay sterile pipettes on the lab bench top. After pipettes are used they should be placed directly into the plastic discard container on the lab bench. In order to make the calculation of the CFU/ml in the original samples less formidable, dilutions are designed to be easy to handle mathematically. The most common dilutions are 1/10 (i.e. 1 ml of sample into 9 ml of diluent) and 1/100 (i.e. 1 ml of sample into 99 ml of diluent), but any dilution can be made. Because dilutions are large when counting bacteria, exponents are used for both the dilutions and the final concentration. All final answers of bacterial concentration should be written with two significant figures, 1 decimal place, in proper scientific notation, i.e. 1.5 x108. (Remember: 1x10a = 10a.) See Figures 7.1 and 7.2 for dilution examples and a sample dilution scheme. In this experiment you will also be using Petrifilm™, a product available from the 3M Company. Petrifilm™ contains a dehydrated medium that needs to be inoculated with one full milliliter of sample to rehydrate the medium. The advantages of Petrifilm™ are its convenience, ease of use, small size, and stability. The disadvantages of Petrifilm™ are its cost, its restriction to a 1 ml sample volume, and the lack of significant colony morphology. There are several different varieties of Petrifilm™ available, each designed to grow a particular organism or group of organisms. We will be using Aerobic Plate Count (APC) Petrifilm™, designed as an all-purpose medium for bacteria, or Coliform Count (CC) Petrifilm™, which is specific for the growth of coliforms, such as E. coli. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Prepare at Home. This will be graded before this experiment is performed. Prepare a neat and easy-to-follow flow chart of your dilutions and platings using information from the following procedure section. This flow chart must be done in the style of Figure 7.1: Sample Dilution Scheme. Indicate which dilutions are prepared, the platings from the proper dilution tubes, and the final dilution on each plate (use Figure 7.2: Determining Dilution for examples). In addition, indicate the total number of pipettes required for this dilution series and indicate on your scheme where you will use each pipette. 5 pts. Figure 7.1: Sample Dilution Scheme

P1

P2

P3

P4

1 ml

1 ml

1 ml

1 ml culture

Legend dilution blank plate PF Petrifilm P

10-3

10-5

9.0 ml 1/10 10-6

P3

P4

P5

99 ml 1/100

9.0 ml 1/10 10-1

P2

99 ml 1/100

1 ml

1 ml 0.1 ml

pipette

0.1 ml

10-3

10

0.1 ml 0.1 ml

-5

10

-2

10

10

-6

1.0 ml

PF

10-7

-4

10-6

Note: if a full milliliter is plated, the dilution on the plate is the same as the dilution of the bottle that the sample came from. If 0.1 ml is plated, the dilution on the plate is 1/10 the dilution of the bottle that the sample came from. Because the standard for microbiology is 1 ml, if any volume less than that is plated it is treated as if it were an additional dilution. Figure 7.2: Determining Dilution Dilution =

_____Volume transferred__________ = Vol. transferred + diluent volume

Volume transferred Total Volume

Ex. #1: 1 ml of culture was transferred to 99 ml of diluent. What is the dilution? Answer: Dilution = 1 ml = 1 ml + 99

1 ml = 100 ml

1 100

or 0.01 or 10-2

Ex. #2: 1 ml of a culture is transferred to 9.0 ml of diluent. From this initial dilution four serial 10fold dilutions are made. Determine the final dilution. Note: to arrive at a final dilution, multiply successive dilutions together. #1 #2 #3 #4 #5 1 ml → 1 ml → 1 ml → 1 ml → 1 ml 9 + 1 ml 9 + 1 ml 9 + 1 ml 9 + 1 ml 9 + 1 ml 1/10 (10-1)

1/10 X 1/10= 1/100 (10-2)

1/10 X 1/10 X 1/10 = 1/1000

1/10 X 1/10 X 1/10 X 1/10 = 1/10,000 (10-4)

(10-3)

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1/10 X 1/10 X 1/10 X 1/10 X 1/10 = 1/100,000 (10-5)

Materials/Organisms per Student E. coli broth culture 4 x plate count agar (PCA) 2 x 9.9 ml dilution blank (silver cap) 4 x 9 ml dilution blanks (blue cap) 7 x 1.0 ml pipettes, sterile 1 Petrifilm™ Petrifilm™ spreader Procedure - Work individually 1. Transfer 0.1 ml of the E. coli culture into a 9.9 ml dilution blank (a tube with 9.9 ml of diluent in it). This is dilution #1. Mix using a vortexer. 2. Using a new pipette, remove 0.1 ml of dilution #1 and transfer into a 9.9 ml dilution blank. This is dilution #2. Mix as above. 3. Using a new pipette, remove 1.0 ml of dilution #2 and transfer into a 9.0 ml dilution tube (dilution #3). Mix as above. 4. Using a new pipette, remove 1.0 ml of dilution #3 and transfer into a fresh 9.0 ml dilution blank (dilution #4) and mix. Using the same pipette, dispense 0.1 ml of dilution #3 onto PCA as well. Spread with a hockey stick. Label the plate with its dilution (as well as your identifying information). 5. Using a new pipette, remove 1.0 ml of dilution #4 and transfer into a fresh 9.0 ml dilution blank (dilution #5) and mix. Using the same pipette, dispense 0.1 ml of dilution #4 onto PCA as well. Spread with a hockey stick. Label the plate with its dilution (as well as your identifying information). 6. Using a new pipette, remove 1.0 ml of dilution #5 and transfer into a fresh 9.0 ml dilution blank (dilution #6) and mix. Using the same pipette, dispense 0.1 ml of dilution #5 onto PCA as well. Spread with a hockey stick. Label the plate with its dilution (as well as your identifying information). 7. Using a new pipette, remove 0.1 ml of dilution #6 and plate onto PCA. At this point, you should have inoculated 4 PCA plates. Also plate 1 ml of dilution #6 onto a Petrifilm™, by lifting up the top cover, depositing 1 ml of sample, dropping the top cover onto the sample, and then spreading it with a plastic Petrifilm™ spreader (this will be demonstrated). Label all plates with their final dilution (as well as your identifying information). Allow Petrifilm™ to solidify for 1 minute before picking up. 8. Rubber band the 4 PCA plates together, invert, and slip in the Petrifilm™ on top. Place in the appropriate incubation tub. All PCA plates and Petrifilm™ will be incubated at 37°C for 48 h. 9. After incubation, count the colonies and record the colony count for each plate/Petrifilm™. Use a VWR pen to mark the colonies on the plate/Petrifilm™ as you count them. Record colony appearance for the Petrifilm™. 10. Determine cell concentration as CFU/ml by averaging the values from all plates/Petrifilm™ that are “countable” (have 25-250 colonies on them). See Figure 7.3 for examples. Record your cell concentration on the class data sheet at the front of your lab bench. Before leaving, record the CFU/ml values for all the other students at your lab bench. Have your TA initial your table.

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Lab Notebook  Describe the appearance of the colonies on the Petrifilm™. Note: the colony characteristics described in Table 6.1 do not apply to colonies on Petrifilm™. 1 pt.  Count the colonies on all of your plates (including the Petrifilm™) and record results in a neatly labeled table. Record results for every dilution - if the plate has more than 250, record as TNTC (“Too Numerous To Count”). 2 pts.  Determine the average number of CFU/ml in your original E. coli culture by averaging all CFU/ml from plates and Petrifilm™ having 25-250 colonies (unless you only have one countable plate). Remember that the exponent should be positive after you finish the calculation. Show your calculations. 2 pts.  Enter your average CFU/ml onto the data sheet at the front of your lab bench. Remember to use only 2 significant figures (one decimal point), exponential form. Copy down the class data from your lab bench data sheet into a neatly labeled table in your lab notebook. 2 pts.  Use the class data from your table to determine the average CFU/ml of the E. coli culture. Show your calculations. 2 pts.  Compare your CFU/ml to the average CFU/ml for your lab bench. Comment on your precision. If there is a discrepancy that is 1 log or greater then provide a possible explanation. 2 pts. Remember that all final answers should be written with two significant figures, 1 decimal place, in proper scientific notation, i.e. 1.5 x108. Use Figure 7.3: Calculating CFU/ml to help set up your calculations. Figure 7.3: Calculating Colony Forming Units (CFU)/ml CFU/ml =

______colony count_______ tube dilution x volume plated

Note: if you’ve determined the final dilution for the plate then volume plated has already been factored in. Just divide colony count by the final dilution. -------------------------------------------------------------------------------------------------------------------Ex: #1: After plating 0.1 ml of a 10-5 dilution, 158 colonies grow. What is the CFU/ml? CFU/ml =

___158___ 10-5 x 0.1

=

_158___ 10-6

6 8 = 158 x 10 = 1.6 x 10

Note: 1/10-6 = 106 (the exponent simply becomes positive) --------------------------------------------------------------------------------------------------------------------Ex: #2: After plating 0.2 ml of a 10-5 tube dilution, 183 colonies grow. What is the CFU/ml? 6 7 ___183___ = _183___ = 91.5 x 10 = 9.2 x 10 10-5 x 0.2 2 x 10-6 ----------------------------------------------------------------------------------------------------------------------Ex: #3: After plating 0.1 ml of a 10-5 tube dilution in duplicate and 0.1 ml of a 10-6 tube dilution in duplicate, you obtain 260 & 245 colonies and 28 & 26 colonies respectively. Determine the CFU/ml.

CFU/ml =

Average CFU/ml =

245 0.1 X 10-5

+

28 0.1 X 10-6 3

+

26______ 0.1 X 10-6

Average CFU/ml = 26.1 x 107 = 2.6 x 108 Note: 260 = Too Numerous To Count or TNTC (this plate count is not used for the calculation)

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EXERCISE 8. SCIENTIFIC WRITING One of the primary objectives of this course is to train you to think in a scientific and critical manner. This means that you will be able to:  develop reasonable hypotheses based on observation  plan precise experiments to test your hypothesis  analyze data and reach valid conclusions In addition, you will be expected to clearly present your scientific work in written communication. The following assignment (see next page) has been designed to help you become familiar with professional scientific literature to see how scientists propose hypothesis, perform experiments, analyze data to reach valid conclusions, and then present their results to the scientific community. Journal Articles. Scientific information in journal articles is normally divided into the following sections: 1. The title of the paper should be brief but must clearly and sufficiently reflect its contents. The title may state the subject of the article or it may give the article's major conclusion. 2. The abstract/summary is informative. It must allow the reader to understand the essence of the authors' research without having to refer to the article. Specific details of data are given, however, methodology is not described in detail unless it is unique. An abstract should be brief but contain the major conclusions. 3. The article begins with an introduction that includes a brief background of the relevant published literature. It usually ends with a statement of the hypothesis the research was designed to solve. 4. A section on materials and methods should be written in enough detail to allow another investigator to duplicate the experiments. New methods are described completely and sources of unique chemicals and equipment are stated. Standard methodologies (e.g., Gram stain, plate count) are not explained. Methods previously described are not repeated, but a reference is given to where they are described in detail. 5. Results are presented in a sequence that logically supports or rejects the hypothesis. Illustrations and tables that accurately reflect the data are included in results. Illustrations and tables should be accompanied by a title and an informative legend. Extensive interpretation of data is not given here. 6. A discussion of the results and the conclusions drawn from the data includes interpretation of its meaning. It should also address any discrepancies between these results and other papers. The potential implications of the work should be stated. 7. References that are cited in the text are listed in a style dictated by the journal. In this class, we will use the style of the American Society for Microbiology (ASM) Journals. When references are cited within the paper, only a number is used. References listed in the References section are numbered in the order in which they appear in the article (citation-sequence reference system). Details on ASM style: citations are arranged in the order they appear in the article. All authors are listed by last name, first initial (i.e. do not use “et al.”) and put in bold type. Author names are followed by the year the article was published, and then the complete article title. The journal name is after the title, abbreviated according to an acceptable style. Volume number is bolded, followed by issue number (if present) in parentheses, colon, page span (starting page – ending page). The page reference includes all inclusive pages. For more information about the ASM reference style, go to: http://jcm.asm.org/site/misc/journal-ita_org.xhtml#02. Examples: Journal article: 1. Lowry J, Hanson J. 1965. Electron microscope studies of bacterial flagella. J. Mol. Biol. 11:293-313. Book: 2. Prescott L, Harley J , Klein L.. 1993. Microbiology Vol. 1, 2nd ed. Wm. C. Brown Publishers, p. 17-25.

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Ex. 8 Assignment - Due Week 3 (10 pts) Note: this is a separate assignment, to be done outside of the laboratory. Your write-up should be typewritten and separate from your lab notebook. (-20% if not typed) Access the following paper at:

http://mbio.asm.org/content/8/3/e00353-17.full or the PDF online at: http://mbio.asm.org/content/8/3/e00353-17.full.pdf+html and then answer the questions below using the accessed paper. The paper will also be available on the MB 303 Canvas site. -------------------------------------------------------------------------------------------------------------------------Article: The Fitness of Pseudomonas aeruginosa Quorum Sensing Signal Cheats Is Influenced by the Diffusivity of the Environment Authors: Anne Mund, Stepher P. Diggle, Freya Harrison doi: 10.1128/mBio.00353-17 2 May 2017 mBio vol. 8 no. 3 e00353-17 Answer the following questions, in your own words: Questions: 1. Define quorum sensing in your own words. 1 pt. 2. What are the lasI and lasR mutants? Why were they used in these research experiments? 1 pt 3. Figure 1 shows the change in Colony Forming Unit (CFU) count as a function of increasing percentages of adenosine. Describe the impact that adenosine as a carbon source has on the three cultures tested and why concentration of adenosine would have different impacts on the three different cultures tested. 1 pt 4. The experimenters then wanted to see how a mixed population of wild type and lasI- mutants would fare with increasing concentrations of adenosine in solid vs liquid medias. a. Print off figure 3 and attach this figure to your written assignment. 1 pt b. What do you observe about the impact on relative fitness that the pure lasI- culture has compared with the mixed culture? 1 pt c. What do you observe about the impact on relative fitness that the cultures in liquid compared to solid? Why is this variable significant? 1 pt 5. Using a database from the OSU Library (such as MedLine), find 2 additional articles on quorum sensing. Do not use any of the articles cited in the assigned paper. Cite the journal articles using ASM style (see details at the bottom of the previous page). Full citation information for the articles must be included. If you cannot find the citation information (i.e. you can only find a reference MedLine number), you should select a different article! 4 pts

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EXERCISE 9. CULTURING THE UNCULTURABLE Introduction Most microorganisms in nature have not been cultured in a laboratory. It is estimated that only 1% of the microorganisms on Earth have been cultured. We do have some ways to get these organisms to grow even if it is not in a “medium” per se or in pure culture. You already studied one when we used hay infusion broth in Exercise 1. Most of the organisms found in hay infusion broth are not easily cultured in pure culture in a laboratory medium. Another way to grow these types of microorganism are with a Winogradsky column. These columns are named after the Russian microbiologist, Sergei Winogradsky, and are a model ecosystem used to study soil and sediment microbiology. Most of the microorganisms found in the columns cannot be grown on ordinary laboratory medium; they are unculturable. In these columns, a core of soil or sediment is mixed with Na2SO4, Na2CO3 and cellulose plus water. It is incubated in the light to allow the growth of photosynthetic organisms. A series of reactions occur in the column as they mature and particular microbial communities develop in specific microenvironments within the column. The height of the column allows an aerobic zone at the surface with microaerophilic and anaerobic zones below. Specific microbial populations grow at different levels because of different environmental conditions. Cellulose is degraded to fermentation products by the anaerobic genus Clostridium. Desulfovibrio uses the fermentation products as an energy source and sulfate as a final electron acceptor and produces hydrogen sulfide that will produce a black color. H2S diffuses to the aerobic zone where it is used by chemolithotrophs such as Beggiotoa and Thiobacillus as an energy source. Thiobacillus can also oxidize ferrous ion to ferric ion and produce ferric oxide (a rust color). Green algae and Cyanobacteria undergo oxygenic photosynthesis producing oxygen for aerobic organisms, while purple and green photosynthetic bacteria undergo anoxygenic photosynthesis. These are just some of the many reactions that result in nutrient cycling in these columns. The Winogradsky columns you will observe were started more than 20 years ago and they have been provided with nothing except water and sunlight since that time. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms Winogradsky columns Procedure – Work individually 1. Observe one of the Winogradsky columns and describe in detail the different layers and colors observed. 2. Record at least 2 organisms seen in the video scope sample, taken from a mixture of the Winogradsky columns. The sample shown on the video scope is a wet mount magnified at 400X. Lab Notebook  Record the different layers and colors of a specific Winogradsky column. 2 pts.  Identify the specific Winogradsky column used by date established and location. 1 pt.  Describe the metabolic/chemical reactions in detail and identify possible organisms responsible for two of the colors/layers observed. See the MB 302 complete textbook for additional information about reactions and possible organisms or look up information online. You must use a source beyond the MB 303 lab manual! 4 pts.  Provide a complete reference for any source of information, in ASM style. 1 pt.  Record at least two different microorganisms you observe on the video scope (400X total magnification). Label in the same manner as Exercise 1, with as much information as possible. Note: the presumptive identification is not required. 2 pts.

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EXERCISE 10. CULTURING BACTERIA: NUTRITION AND ENVIRONMENT Introduction All living organisms require the same elements to grow and divide. CHONPS are needed in macro quantities, K, Mg, Ca, Na and Fe are needed in micro quantities and a number of additional elements are needed in trace amounts. Organisms also require an energy source. The form in which different organisms obtain these elements and energy varies. Some bacteria require organic forms of carbon for both energy and carbon source (chemoorganoheterotrophs). Some bacteria use light energy in photosynthesis (phototrophs) and obtain their carbon from an inorganic source, CO2 (autotrophs). Some bacteria use light energy and organic forms of carbon (photoorganoheterotrophs). Chemolithoautotrophs obtain their energy from inorganic compounds such as H2, NH3, or H2S and use CO2 as a carbon source. See your MB 302 textbook for further details. Most of the bacteria used for the MB 303 lab are chemoorganoheterotrophs. There are many different media used for culturing these bacteria. A medium can be described as all-purpose, meaning it contains all the general ingredients required for growth of non-fastidious microbes and it doesn’t specifically exclude growth in any way. Alternatively, a medium can also be selective, where it contains particular ingredients (salts, dyes, etc), that are used to prevent the growth of specific microbes or it simply lacks ingredients that most microbes require. Defined media are composed of designated amounts of specific compounds; the exact contents are known, down to individual elements. Complex media contain nutrient-rich substances such as yeast extract, peptone, or tryptone and therefore the precise chemical constituents are unknown and will vary slightly from batch to batch. Media can also be differential where the appearance of the microbes on the media will vary, depending upon their interaction with specific ingredients in the media. The difference can either be in colony color or a change in the medium itself, such as a color change or clearing. This is usually achieved with pH indicators that change color with different pHs. Media that lacks any specialized ingredients to distinguish different microbes is called non-differential. To be useful for culturing a particular microbe, a medium must supply the microbe with its basic requirements for growth and lack any inhibitory substances. The basic requirements for growth include 1) water, 2) sources of energy, 3) sources of carbon, nitrogen, and other essential elements, 4) required growth factors (organic compounds such as amino acids or vitamins that the microbe cannot synthesize on its own) and 5) the correct physical environment (temperature, pH, osmotic conditions, oxygen concentration etc). In this experiment we will work with 6 different bacteria, plating them on different types of media and under different environmental conditions, to determine their preferences. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms per Pair 4 plates medium #1 1 plate each of media #2, and #3 1 tube of medium #4 One of the following cultures: Escherichia coli Staphylococcus aureus Micrococcus luteus Bacillus subtilis Enterobacter aerogenes Pseudomonas aeruginosa Procedure – Work in pairs 1. Each pair of students at a lab bench should inoculate the different media with one of the organisms listed above. Plates should be streaked for isolation as in Ex. 6. 2. Four plates are marked "1." They contain the same medium. Each pair of students should use one of the six organisms listed to streak all four plates labeled 1. Streak for isolation on each plate. Label each plate with incubation temperature and conditions (if appropriate), in addition to name/initials, seat numbers, and organism: 37°C for 48 h 37°C for 48 h, anaerobic conditions 50°C for 48 h 13°C for 1 week The anaerobic plates will be incubated in an anaerobic jar. A chemical will be used to remove the oxygen in the chamber, creating an anaerobic environment. 3. Media 2 and 3 will be streaked with the same organism and incubated at 37°C for 48 h. 4. Medium 4 is a broth. Inoculate the broth with a loop, being careful to avoid the inverted Durham tube in the medium, which is used to capture any gas produced by the organism during incubation. Do not markedly tip the tube at any time, as this will introduce a bubble into the Durham tube. This medium will be incubated at 37oC for 48 h. 5. Record the results of all six organisms on all media in the table on the next page. Some results will be covered as a class, with your instructor identifying each medium (media recipes are listed in Appendix B, a glossary of media ingredients is listed in Appendix C). The completed table should be stapled into your lab book. Lab Notebook  Record the observations for all six organisms and the different media/environmental conditions into the table on the following page. Be sure to include information on amount of growth, color of growth, and color of medium, if changed. Amount of growth can be indicated by the legend listed on the table page: growth = (+), heavy growth = (++), slight growth = (+/-), no growth = (-). Staple the completed table into your lab notebook. 6 pts.  Clearly identify your table with an appropriate title and a completed legend, indicating the identity of all media. 2 pts.  Be sure to include the results of all different organisms in the table, as well as temperature of incubation and time of incubation in the column header for each medium. 1 pt.  Indicate on the chart the specific bacterium that you and your partner inoculated by circling that organism. 1 pt.

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Organism

Escherichia coli

Staphylococcus aureus

Enterobacter aerogenes

Bacillus subtilis

Micrococcus luteus

Pseudomonas aeruginosa

#1- 37oC

#1- 50oC

#1- 13oC

#1- 37oC anaerobic #2

#3

#4

Legend:

Media Identification 1– 2– 3– 4–

Amount of Growth - no growth +/- slight growth + growth ++ heavy growth

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EXERCISE 11. INTRODUCTION TO THE SPECTROPHOTOMETER Introduction A spectrophotometer is an instrument used for measuring the effective transmission of monochromatic light through a sample liquid. A beam of light is passed through a suspension and the amount of light transmitted by the suspension is measured. The amount of light transmitted is inversely proportional to the amount of light absorbing or scattering particles in the solution. Before a sample can be measured the instrument must be set to zero with a blank. A blank has all the ingredients of the sample minus the aspect of the sample that you are trying to measure. A spectrophotometer contains a source of white light and an optical system which separates this light into its component wavelengths, collectively called its spectrum. Any wavelength in this spectrum may be selected for use by the operator. The selected wavelength passes through the sample (contained in a cuvette made of optical glass) and strikes a photo-sensitive vacuum tube. The resulting electronic signal is amplified and displayed on the meter, indicating the percent transmittance (how much light passes through the sample) or absorbance (how much light is absorbed by the sample or, in the case of bacteria, how much light is scattered). The spectrophotometer can be used to determine the concentration of a wide variety of light-absorbing molecules and is widely used for enzyme assays and for determining the mass of bacteria in a culture. The theory behind the use of these instruments is related to the Beer-Lambert Law of Solutes, which states that there is a straight line relationship between absorbance (also called optical density or O.D.) of a solution and the concentration of a solute. The optical density is a function of the original intensity of the light beam and the intensity of the beam passing through the solution. You will find that at very high or low solute concentration, the relationship will no longer be linear. This is because a spectrophotometer is no longer sufficiently sensitive when the O.D. is above 1.0 or below 0.05. A bacterial suspension can also be considered the solute (even though it is a suspension of particles), because of its light scattering properties. Therefore a linear relationship exists between its optical density and the number of cells per milliliter. However, the optical density does not directly indicate the number of cells in the population but only the light scattered by the population. To obtain an approximation of population size of a bacterial culture, one must first determine the correct wavelength for the cells and then the O.D. values must be calibrated to a viable cell count to express microorganisms as CFU/ml. This will be performed in Exercise 12: Growth Curve of Escherichia coli. Exercise 11 will allow students the opportunity to familiarize themselves with the spectrophotometer, using a methylene blue solution to mimic a bacterial culture. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials per Pair Spectronic 21 10 ml methylene blue (0.005 mg/ml) 5 pipettes (nonsterile) – 4 x 5 ml; 1 x1 ml 2 spectrophotometer tubes Culture tubes (nonsterile) - to make dilutions Kimwipes Figure 11.1. Spectronic 21 Spectrophotometer. 1. 2. 3. 4.

Meter Pilot Light Wavelength Scale Wavelength Control

5. Power Switch 6. Zero Control for 100% Transmission 7. Sample Holder

2 1

SPECTRONIC 21

7

4 3

6

5

How to Perform Sample Measurements 1. POWER - Flip power switch to on, let warm up for 5 minutes. 2. WAVELENGTH SELECTION - Adjust Wavelength Control to the desired wavelength setting. 3. BLANKING FOR MEASUREMENT* a) Fill a spectrophotometer (spec) tube about ¾ full with distilled water. Water is used as a blank for this experiment since the methylene blue was dissolved in water. You need at least 3 ml of solution in the spec tube for the machine to work properly. b) Tubes should be wiped with Kimwipes before inserting into the Spectronic 21. Insert spec tube into the sample holder, aligning the mark of the test tube with the line on the sample holder. Close the cover. c) Adjust A/T knob (Fig. 11.1, #6) until the meter reads at zero absorbance or 100% transmittance (look at the scale on the bottom of the meter – the meter needle should be all the way to the right). The machine is now ready to measure a sample. 4. SAMPLE MEASUREMENT a) Fill a second spec tube with at least 3 ml of the sample to be measured. b) Remove the blank and replace with the sample. Align the test tube in the same manner as the blank. Close the cover. Record the absorbance (not transmittance) of each sample from the meter. Absorbance is read on the lower scale of the meter. *NOTE: it is necessary to re-blank the machine every time a different wavelength is used. When operating with a fixed wavelength it is necessary to re-blank every 15 minutes to correct for “drift”.

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Procedure - Work in pairs Note: one spectrophotometer tube should be designated as a blank for the entire experiment, to contain nothing but water. Part I Determining the absorption spectrum of methylene blue. 1. This will allow you to choose the wavelength where the absorbance is maximal. The wavelength of maximum absorbance will be used in Part II. 2. Set the wavelength to 675 nm, blank the spectrophotometer with water, and read absorbance (optical density or O.D.) of the methylene blue solution. Note that this is the bottom scale. 3. Repeat Step 2 for the wavelengths 650, 625, 600, 575, 550, 525, 500, 475, and 450 nm. Be sure to reblank with water each time wavelength is changed. 4. From your readings, determine the optimal wavelength. This is the wavelength where you measured the highest optical density. 5. Use a computer program to graph the wavelength (x-axis) vs. O.D. (y-axis), to obtain the absorption spectrum of methylene blue. Staple the completed graph into your lab notebook. Part II The Beer-Lambert Law of Solutes in action. 1. Use water as a blank to adjust the Spectronic 21/20D to 0 absorbance, using the optimum wavelength. 2. Prepare methylene blue dilutions in the nonsterile culture tubes, using the following protocol: dilute the original methylene blue solution 1:2 (add 3 ml of methylene blue solution to 3 ml of water), 1:4 (use 3 ml of 1:2 dilution + 3 ml of water), 1:8 (use 3 ml of 1:4 dilution + 3 ml of water), and 1:10 (use 0.5 ml of original methylene blue solution to 4.5 ml of water). Use the spec tube you used to read the original methylene blue solution but rinse it out with water after each dilution reading. 3. Measure the absorbance of all 5 methylene blue solutions (four dilutions plus the original) at the optimum wavelength determined from Part I. 4. The molecular weight of methylene blue = 320 g/mole. The concentration of the original sample is 0.005 mg/ml. Determine the molarity (M) of your original sample and each dilution. 5. Use a computer program to graph the 5 methylene blue concentrations (including the original sample) in molarity (x-axis) vs. O.D. (y-axis), as a scatterplot or XY graph (with or without lines linking the data points). Add in a trendline. Staple the completed graph into your lab notebook. Lab Notebook  Record the O.D. measurements obtained for Part I and II in neatly labeled tables. 4 pts.  Record the calculated molarities for the original methylene blue solution and the four dilutions. Show your calculations. 5 pts.  Staple in the two graphs for 1) wavelength vs. O.D., and 2) molarity vs. O.D. Make sure that each graph is clearly labeled with a descriptive title, and that the axes are labeled. 8 pts.  Explain what you learned from the Part I graph. 1 pt.  Does the graph from Part II follow the Beer Lambert Law of Solutes? Explain why or why not. 2 pts.

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EXERCISE 12. GROWTH CURVE OF ESCHERICHIA COLI Introduction If a microorganism is inoculated into a flask of medium and the growth rate of this organism reproducing by binary fission is plotted as the logarithm of cell number versus incubation time, a growth curve consisting of four distinct phases results: 1. The initial phase, lag phase, refers to the fact that cell division does not occur immediately; the microorganisms must adjust to their new medium. The length of this stage will depend on the condition of the cells at the time of inoculation and the type of medium into which they were inoculated. If a young culture is transferred into the same medium under the same conditions, no lag time is seen. 2. During the next phase, the exponential phase, microorganisms are growing and dividing at the maximal rate possible given their genetic potential, the nature of the medium and environmental conditions. The population is most uniform in terms of chemical and physiological properties. During this stage the generation time (doubling time) can be determined directly from a graph of log cell number versus time. See the MB 302 textbook for more details. 3. In the stationary phase, growth ceases. Normally this occurs once the population level has reached 109 cells/ml. This stage is reached because available nutrients are depleted, toxic waste products have accumulated, physical space is limited, and/or quorum sensing has occurred. 4. If incubation continues, the culture will enter a fourth phase, the death or decline phase, whereby the number of viable cells decreases exponentially. Growth curves are typically determined using the viable plate method. This method employs spreading a diluted sample of bacteria over a solid medium and determining the number of colonies that arise (see Ex. 7). The number of viable microorganisms in the sample is calculated from the number of colonies formed multiplied by the dilution factor (inverse of the dilution). This method works well as long as microbes are well dispersed so that you increase the chance that each colony arises from an individual cell. On the Petri plates used, this method is only statistically valid if the sample yields between 25-250 colonies. Plates with higher or lower counts are not used normally, although the data should still be recorded. Bacterial cell numbers can also be approximated by measuring cell mass by means of optical density (O.D.) or turbidity (see Ex. 11), once the population has reached about 107 cells/ml. For exponentially growing cells this method is fairly reliable and provides an almost instantaneous estimation of the bacterial concentration for a culture. If cell number versus O.D. are graphed a straight line should result, indicating correlation between the two values. This follows the Beer-Lambert Law of Solutes, which typically applies to molecules in a solution but which has been shown to also apply for bacteria in suspension. A microbiologist can produce a standard curve by measuring both O.D. and cell numbers by plate count for a bacterial population. The resulting graph can be used to extrapolate cell concentration by O.D. in future experiments, thus eliminating the delay caused by incubation. Cell size influences the results obtained by spectrophotometry so the same bacterium must be used and the values are only valid for cells in the exponential phase of growth. In this experiment, we will determine the growth curve of E. coli under various nutritional and environmental conditions. Objectives Lab objectives are not required in the lab notebook for this experiment. The results for this experiment will be written up separately from the lab notebook.

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Materials and Organisms per Group E. coli culture grown overnight 50 ml of brain heart infusion (BHI) medium or 50 ml of minimal salts broth in side arm flask Tube of BHI or minimal salts broth as blank Spectronic 21or 20D with black velvet cloth TSA plates (2 for each sampling time) 1 can of sterile 1 ml pipettes 0.1 % peptone dilution blanks: 9.0 ml (clear liquid, blue cap) 0.1 % peptone dilution blanks: 9.9 ml (clear liquid, silver cap) 1 package of Kimwipes 1 hockey stick, ethanol Procedure - Work in groups of 4-5 Treatment Medium Temp Inhibitor # 1 BHI broth 37oC 2 BHI broth 25oC 3 BHI broth 46oC 4 BHI broth 37oC 50g/ml chloramphenicol 5 Minimal salts broth 37oC

O2

Tape Color

shaking no shaking shaking shaking shaking

orange blue red pink white

Overview: each group will inoculate a flask with a specified amount of an overnight culture of E.coli. Immediately after inoculation, each group will take both an initial O.D. and a viable cell count sample. Additional O.D. measurements will be taken every 15 minutes over a 3 hour period, while additional viable plate count samples will be taken every 30 minutes over a 3 hour period. The dilution schemes are shown in Figure 12.1. For an O.D. measurement at any time point: if the O.D. is below 0.5, use dilution scheme A. If the O.D. is over 0.5, use dilution scheme B. 1.

Blank your spectrophotometer with an appropriate blank (uninoculated BHI or minimal salts broth) at 600 nm. Use the black velvet cloth to prevent light from entering while you are measuring and use Kimwipes to wipe your tubes before measuring. Re-blank before every reading.

2.

Vortex the overnight E. coli culture for a few seconds, to get all the cells in suspension. Immediately inoculate the side-arm flask with the specified amount of culture, for an O.D. value of ~ 0.07. (Note: this is less than 0.5, so every group should start with Dilution Scheme A shown on Figure 12.1)

3.

Immediately after inoculating flask with E. coli, measure and record the time zero O.D. reading of the culture in the flask at 600 nm. Fill out the data chart provided as you take readings. It may take two people to measure the O.D. with the slightly cumbersome side arm flasks. Fill side arm with culture, wipe glass with Kimwipe. Place the arm into the chamber, in the same direction each time you take a measurement. Make sure the side arm is straight upright and snugly fit against the right side of the measurement chamber, otherwise the O.D. will be drastically off. Cover the flask with black velvet cloth during the reading. If you are having trouble, ask TA for help.

4.

After measuring the O.D. at time zero, pour side arm contents back into main flask and then pull sample for the time zero dilution scheme. Remove 1 ml of culture from the flask (using aseptic technique), dispense into a labeled 9.0 ml dilution blank and then set it aside.

5.

Place your flask into the appropriate shaker incubator (if your group is treatment 2, leave the flask on the lab bench at room temperature, no shaking). Be sure to use caution when handling and removing flask from shaker. These flasks break easily and are very expensive. The time the flask is out of the shaker should be kept to a minimum.

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6.

Vortex the dilution tube that you set aside in Step 4 for several seconds. This is to separate and homogenize the bacteria. Finish Dilution Scheme A on Fig. 12.1. Vortex subsequent dilution tubes thoroughly before transferring to next dilution. Make sure to use a new sterile pipette for each dilution. The same pipette can be used to make a dilution and a plate if both samples come from the same dilution tube. Remember to sterilize hockey sticks by dipping in alcohol and flaming. Plates should be spread immediately after sample is deposited onto them. Do NOT allow the plates to sit before spreading them. Be sure to mark each plate with group ID, treatment #, time of plating, and final dilution. Every group should be preparing 2 plates for each sampling time (i.e. every 30 minutes). Once the 2 plates are prepared, it is recommended that used dilution blanks be discarded before the next sampling time, to prevent confusion.

7.

Groups should continue to take O.D. readings of the culture every 15 minutes for 3 hours, filling out the data sheet provided as readings are obtained. Dilutions/platings should be performed every 30 minutes for 3 hours. The OD reading will determine which dilution scheme (A or B) is appropriate. Perform dilution scheme A if the O.D. is less than 0.5. Perform dilution scheme B if the O.D. is at or greater than 0.5.

8.

After the last reading, place a rubber band around all the plates for your group and place in the appropriate incubation tub. The plates will be incubated at 37°C for 48 h. Remove the tape from your side arm flask and carefully place it into a metal discard coffin.

9.

Hand in the group data sheet to your TA with the names & seat #s of all the group members. 2.5 inclass points.

Figure 12.1 Flow chart of dilutions for viable cell count Dilution Scheme A - for O.D. readings less than 0.5: 1.0 ml

0.1 ml

0.1 ml

1.0 ml

9

flask

9.0 ml

9.9 ml

Tube dilution:

10-1

10-3

9.9 ml 10-5 Plate 0.1 ml

Dilution Scheme B - for O.D. readings at 0.5 or above: 0.1 ml 0.1 ml 0.1 ml

Tube dilution:

9.9 ml 10-2

10-4

10-6 Plate 0.1 ml

Plate dilution:

45

10-7

1.0 ml

9.9 ml

9.9 ml

10-6 Plate 0.1 ml

10-6

Plate dilution:

flask

9.0 ml

10-7

9.0 ml 10-7 Plate 0.1 ml 10-8

Next Lab Period (after plates have been incubated)  Count the colonies on the plates that your group prepared. If the count goes above 250 colonies on a single plate, record the colony count for that plate as TNTC. Record the information on the data sheet provided and hand in to your TA at the end of lab. 2.5 in-class points.  A complete set of data for all treatments will be posted on Canvas. Use this information to complete the assignment described on the following pages. Note: it is not required to record anything in your notebook for this experiment. However, each student should make note of the treatment that their group was assigned.

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The Ex. 12 assignment is due on the date specified in the lab syllabus (Week 9) and does not go into your lab notebook. It should be handed in as separate assignment - typewritten, all pages stapled together as one packet, clearly labeled with name & seat #. 40 pts total Access the class data for Ex. 12 on Canvas, in the Course Information module 1. Access the class data on Canvas. Use class data from the treatment for your group and Treatment 1 (if your group did Treatment 1, choose one additional treatment). Place colony counts/ODs in a neatly labeled table. Calculate the average CFU/ml for each time point for the 2 treatments, using appropriate colony counts. Circle or bold the colony counts that were used. Be sure to use correct notation and significant figures (See Ex. 7). Even though you have more than one dilution per time point, all the appropriate data needs to be averaged together so that you end up with only one CFU/ml for each time point. Show a sample calculation for one time point for each treatment. See Figure 12.2 for sample calculation. 10 pts. 2. Using the semi-logarithmic graph paper provided on the following page: a. Graph Time (minutes) vs O.D. for Treatment 1 + one other treatment. b. Graph Time (minutes) vs Log Cell Number per ml for Treatment 1 + one other treatment. (Note: if correct notation and sig figs are used then “Log Cell Number per ml” is the same as CFU/ml. For example, 3.7 x 104 CFU/ml is a log cell number). Use left y-axis for O.D and right y-axis for Log Cell Number per ml as shown in the sample graph (Figure 12.2), with all four curves on the same graph. Each curve must be a different color. Look at the time vs. O.D. curve to determine if a lag and/or stationary phase is present. Using this information, draw in a best-fit line for the exponential phase of the time vs log cell #/ml data only. Use this best-fit line for step #3 below. See Figure 12.3: Sample Graph for how to set up the scale and use as much of the graph paper as possible to separate the O.D. and cell number curves. Start the cell number scale with the lowest exponent value possible (i.e.106) so that the O.D. and cell number graphs are separated. You may do this graph by hand or computer, as long as you know how to do 2 separate y-axes. 11 pts. 3. Determine the generation time directly from the best-fit line for the exponential phase of the Time vs. Log Cell Number/ml curve for Treatment 1. Generation time is the time it takes for a population to double in size. Select 2 points in the exponential growth phase that represents a doubling in the population size. Draw horizontal lines from each point to intersect with the bestfit line. Draw vertical lines down to intersect with the x-axis. Determine the difference between the 2 points. This is the generation time in minutes. Be sure to clearly show how you determined this number from the graph. A number without indication of how it was determined is worth no points. Do not use the generation time calculation. For your 2nd treatment, determine the generation time if you can; if you can’t, explain why you cannot. 5 pts. 4. Use a computer to graph cell concentration as CFU/ml (x-axis) vs. O.D. (y-axis) for the Treatment 1 exponential data points only (as determined by step #2), using a scatter-plot graph format. The CFU/ml may be input as a whole number or exponential form (370000 or 3.7E5) but format the x-axis on a logarithmic scale (click on the x-axis for format options). Format the x-axis minimum so the data points are spread over the entire graph. For example, if the lowest value is 2.0 x 107 CFU/ml, type in 1E7 as a minimum. Add in a best-fit (or trend-line). 3 pts. Explain why exponential data points only were used. How would it skew the data if lag or stationary phase data points were used? 1 pt. Explain what this graph demonstrates about the experiment. 2 pts. 5. Explain how temperature, nutrient level, oxygen level and the antibiotic chloramphenicol affect bacteria by interpreting the results obtained for treatments 2-5. It is not sufficient to say that the condition slows or speeds up growth, but clearly indicate why each condition would decrease or increase growth. What impact does each condition have on bacterial cells, physiologically? 8 pts.

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Figure 12.2 Sample calculation of CFU/ml for a single time point of growth curve experiment Time OD Dilution # OF COLONIES (min) (actual) 600 nm 30 10-6 TNTC, 254, 240, 268, 251, TNTC 10-7 0, TFTC, 27, 31, 23, 32 Note: since only those plates that have between 25-250 colonies are considered statistically significant, only 4 colony counts will be used for calculation of the average CFU/ml (numbers in bold).

Here are two ways to obtain an average CFU/ml for this time point: 1. Add all the colony counts together, dividing each by their respective dilution. Then divide by the number of statistically significant plates, to get an average CFU/ml: 240 + 27 + 31 + 32 =114 x 107 = 2.9 x 108 CFU/ml -6 -7 -7 -7 10 10 10 10 4 2. Convert all the plate count values to the same exponent. For example, 240/10-6 is the same as 24/10-7. (Note: this does not change the fact that the original colony count, 240, was statistically significant). Add all the statistically significant values, divide by the total, and multiply by the appropriate exponent (be sure to make it positive to reflect the dilution factor, not the dilution): 24 + 27 + 31 + 32 x 107 = 2.9 x 108 CFU/ml 4 Same answer, just two different ways of setting up the problem. Figure 12.3 Sample graph for Step #2 (one treatment only)

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108

Time (minutes)

Log Cell Number per ml

O.D. 600 nm

1010

▲= O.D. 600 nm = log cell number/ml Line = best fit line for log phase

Note: student graphs should contain 4 curves, to represent 2 different treatments (Treatment 1 & one additional treatment). Different colors must be used to highlight the different curves.

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EXERCISE 13. IDENTIFICATION OF UNKNOWNS Introduction For the next four laboratory periods each student will work on identifying two unknown bacteria. See Figure 13.1 for a list of the possible bacteria and their major characteristics. Students will perform standard tests to identify bacteria in exercises 14-16. Tests performed in Ex. 14 are designed to separate major divisions while tests performed in Ex. 15 and 16 are designed to separate more closely related organisms. One of the first things that each student will do is prepare two flowcharts from the information contained in Table 13.1, one for the gram positive bacteria and one for the gram negative bacteria. The flowchart will be in the form of a graphical dichotomous key (see Appendix E-1 for example). A flowchart is not designed to encompass all information known about each organism, rather it is designed to help streamline the preliminary identification of an organism. Clinical microbiologists often rely upon flowcharts to help guide their initial testing, in order to save time, energy, and supplies. Flowchart construction for each set of unknown bacteria will provide students with a better understanding of the tests to be used, as well as information regarding the reliability of each test. MB 303 flowcharts should separate out all eight bacterial possibilities for each Gram reaction, following the detailed instructions listed on the next page. Some students will be able to identify their unknown after one or two tests. However, the purpose of this exercise is not only to identify the bacteria but also to check aseptic technique and ability to accurately record and interpret observations. You will be required to transfer your two unknown bacteria numerous times and into numerous media without contaminating or switching the two! Carefully observe each unknown on the slant when you receive it and record how it looks. If the appearance changes as you proceed, check with the instructor. Students will receive their unknowns on a Brain Heart Infusion (BHI) agar slant. The BHI slant is your working stock culture. You will need to subculture to a new slant at each lab period, as it is vital to have a fresh culture for setting up various biochemical tests. You will perform various biochemical tests on the unknowns at the same time as you are doing these tests on known bacteria. You should always observe and interpret the tests inoculated with the known bacteria first, so that you have a control for comparison to your unknowns. That way you will know exactly what a positive reaction looks like versus a negative reaction. The tests will allow you to determine various characteristics about the unknowns, allowing for their identification. Keep a detailed record in your lab notebook of all tests performed, observations of test results, and interpretation of the results. An observation is what you observe (i.e what color did it change and at what point?), while an interpretation explains the observation (“the medium changed to pink after the addition of chemical X because the organism has the enzyme Y”). Objectives Lab objectives are not required in the lab notebook for this experiment

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Materials per Student Lab #1: Unknown A, Unknown B (on BHI slants) 4 BHI slants (slant, dark blue cap) Note: requests for Lab #2: 2 BHI slants additional media due 2 motility agars (semisolid deep, olive cap) 2 TSA plates (dark pink stripe on side) to student error will 2 nitrate broths (yellowish broth, black cap) result in a point 2 OF deeps (green deep, white cap) deduction. 1 tube mineral oil (clear viscous liquid, clear cap) Lab #3: 2 BHI slants 1 mannitol salts agar (MSA) plate (pink agar) 2 lactose broths (purple broth, green cap) 1 ornithine decarboxylase deep (purple deep, purple cap) 1 urea broth (orangish broth, red cap) 1 tryptone broth (clear broth, turquoise cap) 2 MRVP broths (yellowish broth, brown cap) 1 citrate slant (green slant, yellow cap) Lab #4: 2 TSA plates Procedure – Work individually Lab #1 1. Perform a Gram stain on each unknown. Determine which organism is gram positive and which is gram negative. Show each Gram stain to your TA (2.5 pts total). If the TA asks you to redo your Gram stain, it is because they want to be sure that you have the correct observation/organism. Record detailed observations about initial BHI growth and gram stain results in lab notebook. 2. After TA verification, transfer both unknowns to 2 BHI slants. Incubate one slant of each unknown at 37°C and another slant of each unknown at 30oC. From now on always label cultures as gram positive or gram negative (opposed to “A” or “B”), to prevent confusion. Preparation for Lab #2 1. Prepare two flowcharts from information given on the possible unknowns (see Table 13.1), one for the 8 possible gram negative bacteria and one for the 8 possible gram positive bacteria. Each flowchart must be a graphical dichotomous key that separates the possible bacteria into smaller and smaller groupings based on major characteristics, eventually separating all possible bacteria from one another (see example flow chart, Appendix E, for proper format) (20 pts). Follow these instructions carefully:  Each flowchart should be typewritten and on a separate sheet of paper (not back to back).  Clearly label each juncture with the medium or test being used. Label each arm with the possible product or result. Be specific about what the medium/test is and what the results could be (i.e. ferments what? Reduces nitrate to what?). Listing “positive/negative” or “yes/no” is not sufficient.  The 1st division for the gram negative bacteria must be the oxidase test (NOT the oxidativefermentative test!).  The 1st division for the gram positive bacteria must be morphology (cocci vs. bacilli).  Select tests that will provide the least ambiguous results (i.e. do not separate 2 organisms based on a positive vs. weakly positive reaction).  Do not use all the characteristics possible, just those that will definitively separate the organisms.  Do not use motility or cell arrangement to divide bacterial possibilities, as these characteristics are dependent on medium used and the incubation environment.  Pigmentation/lack of pigmentation should only be used to separate organisms if pigmentation characteristics are specified. Otherwise it should be assumed that the organism is inconsistent.  Be sure to indicate scientific names correctly (full genus & species, in italics). This is an INDEPENDENT assignment. Flowcharts identical in aspects of style, form, or function will receive no credit. Flowcharts using organisms from previous terms will receive no credit.

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Record all observations/interpretations for unknowns into a table(s) in your lab notebook during lab time. Your notebook will be checked for these observations/interpretations. Be sure to observe the reactions of the known bacteria first, before interpreting your unknowns. Lab # 2 1. Observe your BHI slants. Determine if each unknown grows significantly better or only at one temperature over the other, or if it produces a pigment at only one temperature. Once the optimum temperature of each unknown has been determined (based on growth or pigmentation), always incubate at that temperature. If the unknown grows equally well at both temperatures and does not produce a pigment at only one temperature, select 37oC as the incubation temperature. 2. Transfer a small amount of each unknown to a fresh BHI slant, using a sterile inoculating loop. 3. Inoculate each unknown into motility agar by using a sterile inoculating needle to remove a small amount of the culture off the BHI slant, inserting the needle to the bottom of the deep, and then pulling the needle straight back out. 4. Inoculate each unknown into a nitrate broth tube, using a sterile inoculating loop. 5. Streak each unknown onto a TSA plate, streaking for isolation using the method from Ex. 6. 6. Inoculate your gram negative unknown only into 2 tubes of OF medium, using a sterile inoculating needle. Pour a small quantity of sterile mineral oil over the agar surface of 1 OF tube, about 1 cm deep. This will be the “closed tube.” The tube without mineral oil will be the “open tube.” Note: the gram positive unknown does NOT go into OF tubes; this test is inappropriate for gram positive bacteria. 7. Place all media into the correct incubation tub(s) (based on the selected temperature for each unknown), after appropriately labeling. Lab #3 1. Observe your BHI slants. Transfer each unknown to a fresh BHI slant, using a sterile inoculating loop. Place tubes into the correct incubation tub(s), after appropriately labeling the tubes. 2. Observe the motility agar for each unknown. Determine the motility ability and oxygen requirement of each unknown, using the information on the following page. These are two separate results using this one medium, so be sure to indicate a separate observation and interpretation for each result. 3. Observe the nitrate broth for each unknown for any coloration. Then use the instructions in Ex. 14 to complete the test. 4. Observe and interpret the OF tubes for the gram negative unknown, using instructions in Ex. 14. 5. Observe TSA plate for each unknown, checking purity and colony characteristics (use Table 6.1). 6. Perform an oxidase test on your gram negative unknown, using an isolated colony on the TSA plate. Perform and interpret the test using instructions found in Ex. 14. 7. Perform a catalase test on your gram positive unknown, using an isolated colony on the TSA plate. Perform and interpret the test using instructions found in Ex. 15. 8. Inoculate your gram positive unknown onto a mannitol salts agar (MSA) plate (streaking for isolation), a lactose broth tube, and an MRVP broth tube, using a sterile inoculating loop. 9. Inoculate your gram negative unknown into a ornithine decarboxylate deep, urea broth, tryptone broth, MRVP broth, citrate slant, and lactose broth, using a sterile inoculating needle/loop. 10. Place all media into the correct incubation tub(s), after appropriately labeling. Lab #4 1. Observe and interpret the media inoculated with your gram positive unknown, using the instructions found in Ex. 15. The lactose broth is also described in Ex. 16. 2. Observe and interpret the media inoculated with your gram negative unknown, using the instructions found in Ex. 16. 3. Prepare a single Gram stain composed of a mixture of your unknowns by mixing a small amount of each culture together with a small amount of water. Show to your TA in focus under 100X, with the ideal location selected. Students must perform this independently, with no assistance from other students. Assistance from the instructor or TAs will result in docked points.

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The Gram stain will be graded on appropriate color/morphology, presence of both unknowns, and ability to focus the microscope. Record observations in your lab notebook. 5 pts. 4. Streak each unknown for isolation onto a TSA plate. Label each plate with your full name, unknown number, and if it is the gram + or – unknown. Place in incubation tube for selected temperature. The plates will be graded for purity (2 pts), good streaking technique (1 pt), isolation of at least 1 colony (1 pt), and good isolation of numerous colonies (1 pt). 5 pts each. Motility Agar* Motility agar is an excellent way to determine the motility of most bacteria. It contains one-fifth the agar normally found in solid medium and also contains gelatin. This makes the medium semi-solid and allows the bacteria to move out into the medium if they are motile. Here are the possible observations: 1) Nonmotile bacteria will only grow in the stab line. The stab will not appear 3-dimensional when rotated; often the stab has a “dagger-like” appearance where it’s thicker in one view, and then very thin in another view. This is due to the fact that the needle often moves slightly from side to side during the inoculation of the tube. 2) Motile bacteria will move away from the stab line. In fact, very motile bacteria will make the entire medium turbid. If you are unsure if the entire tube is turbid or not, compare it to a tube of uninoculated motility agar. 3) If the organism only grows on the surface of the medium this test is inconclusive for motility (typically additional tests would be performed in order to determine motility). The interpretation in your lab notebook should state: “motility cannot be determined.” Oxygen Requirement* Bacteria show considerable variation as to their requirement for and tolerance of oxygen. The following are definitions of some of the terms used to distinguish bacteria in regard to their oxygen requirements: 1) Aerobe - an organism that grows in the presence of oxygen and is able to use oxygen as a final electron acceptor in respiration. An aerobe will only be able to grow on the very surface of motility agar, with no growth down in the tube at all. 2) Anaerobe - an organism that can grow in the absence of oxygen. There are different categories of anaerobes: a. Strict or obligate anaerobe - an anaerobe which is killed by the presence of oxygen. This type of anaerobe will only grow below the penetration point of the oxygen, typically the bottom of the tube. b. Aerotolerant anaerobe - an anaerobe which does not utilize oxygen as a final electron acceptor but can grow and tolerate the presence of oxygen. This organism can grow everywhere in a motility tube, with equal amounts of growth throughout. c. Facultative anaerobe - an organism that prefers to grow in the presence of oxygen utilizing oxygen as the final electron acceptor, but in the absence of oxygen, it can ferment or undergo anaerobic respiration. This organism can grow everywhere in a motility tube, but there should be increased amounts of growth where oxygen is present. It can be difficult to distinguish between an aerotolerant anaerobe and a facultative anaerobe. 3) Microaerophiles – an organism that cannot grow in the normal 20% oxygen concentration found in the atmosphere but instead grows in a limited oxygen environment of 2-10%. This organism will form a thin band of growth in a motility tube, a short distance below the agar surface. Typically Gas pac jars are used to culture anaerobic or microaerophilic bacteria in the laboratory or to demonstrate the oxygen requirement of bacteria. Chemicals are used to create these reduced oxygen environments by generating H2 that reacts with the oxygen present in the jar to form H2O. The amount of H2 generated determines whether or not all the O2 will be converted to H2O (obligate anaerobic conditions) or some O2 will remain (microaerophilic conditions). Due to the difficulty in working with these organisms, none of our cultures are microaerophilic or obligate anaerobes. *Note: if you have an organism that makes a diffusible pigment (i.e. one that can travel away from the bacterium), be sure that you look for where growth occurs, not pigmentation. In addition, some organisms can only make a pigment in the presence of oxygen, but can grow elsewhere.

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Procedure The numbers after the tests indicate the exercise in which the test is described. The capital letters after the tests are to indicate the order these tests should be recorded on the final unknown report. All observations for your unknowns should be recorded in your lab notebook. Lab #1. Perform a Gram stain & inoculate 2 BHI slants for each unknown: Gram+ Gram1. Gram stain – Ex. 5 (A) 1. Gram stain – Ex. 5 (A) 2. BHI slant, incubate at 37oC – Ex. 3 (B) 2. BHI slant, incubate at 37oC - Ex. 3 (B) o 3. BHI slant, incubate at 30 C (C) 3. BHI slant, incubate at 30oC (C) Lab # 2. Record results of BHI slants at different temperatures and determine the optimum temperature for each unknown. Inoculate the following: Gram+ Gram1. BHI slant 1. BHI slant 2. Motility agar – Ex. 13 (D, E) 2. Motility agar – Ex. 13 (D, E) (for motility + oxygen requirement) (for motility + oxygen requirement) 3. Nitrate broth – Ex. 14B (F) 3. Nitrate broth – Ex. 14B (F) 4. TSA streak plate – Ex. 6 (G) 4. OF deep (2 tubes) - Ex. 14A (G) 5. TSA streak plate – Ex. 6 (H) Lab #3. Record results of previous tests, perform catalase/oxidase tests, and inoculate the following: Gram+ Gram1. BHI slant 1. BHI slant 2. Catalase (from TSA plate) - Ex. 15A (H) 2. Oxidase (from TSA plate) – Ex. 14C (I) 3. Mannitol salts agar (MSA) – Ex. 10, 15 (I) 3. Ornithine decarboxylase – Ex. 16A (J) 4. MR, VP broth – Ex. 15B (J, K) 4. Urea broth – Ex. 16B (K) 5. Lactose broth – Ex. 15 (L) 5. Tryptone broth – Ex. 16C (L) 6. MR, VP broth – Ex. 16C (M, N) 7. Citrate slant- Ex. 16C (O) 8. Lactose broth – Ex. 16D (P) Lab #4. Record results of previous tests. Perform the following: Gram+ Gram1. Gram stain – show to TA & record results 1. Gram stain – show to TA & record results 2. TSA streak plate 2. TSA streak plate Ex. 13 point breakdown (100 pts total) 1. Lab 1: Gram stain of each unknown. Record results in lab notebook. 2.5 in-class pts. 2. Lab 2: Initial flowcharts of gram positive and gram negative possibilities (see “preparation for lab #2” for directions). Flowcharts must be typewritten (see example, Appendix E). 20 pts. 3. Lab 4: Combined Gram stain, streak plate of each unknown. Record Gram stain results in lab notebook. 15 pts. 4. Final reports must be typewritten and should include (62.5 pts): a. Results Chart. The results chart for each unknown (on separate pieces of paper) must have 3 columns: 1) tests/media used, 2) observations, and 3) interpretations, with organism identified at the bottom and exceptions to the characteristics listed (see examples, Appendix F). Each report should be labeled with student name, unknown number and gram positive/negative unknown. The tests/media used must be listed in the order given on this page. Positive and negative are not acceptable interpretations (i.e. do not say “lactose negative,” say “cannot ferment lactose”). b. Flowchart. Attach the proper flowchart to each unknown report. Indicate on the flowchart how you made your decision by highlighting the path you followed.

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Possible gram positive unknowns

Table 13.1: List of Possible Unknown Bacteria and their Major Characteristics Note: If a characteristic is not listed it means it is not reliable/conclusive for that organism Bacterium Characteristics Bacillus cereus gram+ bacilli (filamentous), motile, facultative anaerobe, reduces nitrate to nitrite, has catalase, MR+, VP-, no growth on mannitol salt agar, lactose nonfermenter Bacillus gram+ bacilli (filamentous), motile, aerobe, does not reduce nitrate, has megaterium catalase, MR+, VP-, no growth on mannitol salt agar, lactose fermenter Bacillus subtilis gram+ bacilli (filamentous), aerobe, reduces nitrate to nitrite, has catalase, MR-, weak VP+, growth on mannitol salt agar/no or slight mannitol fermentation, lactose non-fermenter Enterococcus gram+ cocci in chains, nonmotile, aerotolerant anaerobe, does not reduce faecalis nitrate, lacks catalase, MR+, VP-, growth on mannitol salt agar/mannitol fermentation, lactose fermenter Lactobacillus gram + bacilli, nonmotile, facultative anaerobe, does not reduce nitrate, lacks brevis catalase, MR+, VP-, no growth on mannitol salt agar, lactose fermenter Micrococcus luteus gram+ cocci (tetrads), bright yellow pigmentation, nonmotile, does not reduce nitrate, has catalase, MR-, VP-, growth on mannitol salt agar/no fermentation, lactose non-fermenter Staphylococcus gram+ cocci (clusters), light orange-yellow pigmentation, nonmotile, aureus facultative anaerobe, reduces nitrate to nitrite, has catalase, MR+, VP-, growth on mannitol salt agar/mannitol fermentation, lactose fermenter Staphylococcus gram+ cocci (clusters), nonmotile, facultative anaerobe, no or slight nitrate epidermidis reduction, has catalase, MR+, VP- or weak VP+, growth on mannitol salt agar/no fermentation, lactose fermenter Alcaligenes faecalis

Possible gram negative unknowns

Enterobacter aerogenes Escherichia coli

Klebsiella pneumoniae Proteus vulgaris

Pseudomonas aureofaciens Pseudomonas aeruginosa Serratia marcescens

gram- bacilli, motile, aerobe, has oxidase, oxidative, does not reduce nitrate, lacks ornithine decarboxyalse, lacks urease, lacks tryptophanase, MR-, VP-, has citrase, lactose non-fermenter gram- bacilli, motile, facultative anaerobe, lacks oxidase, oxidative/fermentative, reduces nitrate to nitrite, has ornithine decarboxylase, lacks urease, lacks tryptophanase, MR-, VP+, has citrase, lactose fermenter gram- bacilli, motile, facultative anaerobe, lacks oxidase, oxidative/fermentative, reduces nitrate to nitrite, lacks ornithine decarboxylase, lacks urease, has tryptophanase, MR+, VP-, lacks citrase, lactose fermenter gram- bacilli, facultative anaerobe, lacks oxidase, oxidative/ fermentative, reduces nitrate to nitrite, lacks ornithine decarboxylase, has urease, lacks tryptophanase , MR+, VP-, has citrase, lactose fermenter gram- bacilli, facultative anaerobe, lacks oxidase, oxidative/fermentative, reduces nitrate to nitrite, lacks ornithine decarboxylase, has urease, has tryptophanase, MR+, VP-, lacks citrase, may or may not ferment lactose gram- bacilli, grows at 30o & 37oC but only produces orange pigment at 30oC, has oxidase, oxidative, reduces nitrate to nitrite, lacks urease, lacks tryptophanase, MR-, VP-, has citrase, lactose non-fermenter gram- bacilli, produces blue-green pigment at 30o & 37oC, motile, facultative anaerobe, has oxidase, oxidative, reduces nitrate to nitrogen gas, lacks urease, lacks tryptophanase, MR-, VP-, has citrase gram- bacilli, grows at 30o & 37oC but only produces red pigment at 30oC, motile, facultative anaerobe, lacks oxidase, oxidative/fermentative, reduces nitrate to nitrite, has ornithine decarboxylase, lacks urease, lacks tryptophanase, MR+, VP-, has citrase, lactose non-fermenter

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EXERCISE 14. RESPIRATION/FERMENTATION/ANAEROBIC RESPIRATION Introduction Most chemoorganoheterotrophic bacteria catabolize carbohydrates. Carbohydrates are organic molecules that are composed of carbon, hydrogen, and oxygen in the ratio C(H2O)n. Carbohydrates are divided into classes dependent upon the size of the molecule. Monosaccharides such as glucose are single molecules composed or 4-7 carbons. Disaccharides such as lactose are composed of two monosaccharides. Polysaccharides such as starch are composed of 8 or more monosaccharide units. Most carbohydrates used by microorganisms are converted to glucose. The process by which glucose is metabolized varies with the organism and can also be affected by the conditions under which the organism is cultivated. Fermentative metabolism does not require oxygen. Glucose is degraded by a pathway such as the Embden-Meyerhof pathway. Oxidations occur during this pathway and pyruvate is a common final product. In fermentation no outside final electron acceptors are present so pyruvate also serves as the final electron acceptor, resulting in the formation of organic acids or alcohols. In fermentation, ATP is only formed by substrate level phosphorylation. Respiratory metabolism requires a functional electron transport chain. Examples of electron carriers involved in the electron transport system are NAD, FAD, cytochromes and quinones. The type of carriers can vary. A cytochrome system is essential for the last steps by which oxygen is reduced. The presence of a c-type cytochrome correlates with a positive oxidase test. This test can be used to separate the Enterics (gram-negative bacilli in the family Enterobacteriaceae, that lack the cytochrome c oxidase enzyme) from gram negative non-Enterics such as Alcaligenes, Neisseria, and the Pseudomonads, that have the cytochrome c oxidase enzyme. The oxidase reagent (tetra-methyl-paraphenylene diamine HCl ) acts as an artificial electron donor for the cytochrome c oxidase enzyme, turning a pink-maroon-black color when oxidized. Enterics also have cytochromes but not specifically cytochrome c. The electron transport chain of Enterics is shorter and less ATP is produced. When oxygen is the final electron acceptor, CO2 and H2O are the end products. Oxidative-fermentative media (OF medium) is used to determine whether an organism metabolizes carbohydrate strictly by oxidative respiration, by fermentation, or both. The carbohydrate substrate (typically glucose) is added to a basal media which also contains peptone. Bromthymol blue is the indicator dye. It is green at a pH near neutrality, yellow at an acid pH and blue at an alkaline pH (>7.5). Bromthymol blue also inhibits the growth of most gram positive organisms, making the medium selective and differential. Anaerobic respiration occurs when bacteria use a substance other than oxygen as the final electron acceptor during respiration. A commonly used alternative acceptor is the nitrate ion. Some bacteria reduce nitrate to nitrite in the process. Other bacteria further reduce nitrate to nitrous oxide or nitrogen gas. Bacteria that undergo these reactions typically only use nitrate if oxygen is not available in the environment, since aerobic respiration yields more ATP for the cell. Nitrate broth is used to determine a bacterium's ability to reduce nitrate, using the process of anaerobic respiration. Nitrite is detected by the addition of sulfanilic acid (Nitrate A) and dimethyl-alpha-naphthylamine (Nitrate B) to inoculated nitrate broth, while nitrogen gas is detected by the addition of powdered zinc to inoculated nitrate broth (after reagents A & B have been added). Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms per Pair 1 culture: Klebsiella pneumoniae or Pseudomonas aeruginosa TSA plates of K. pneumoniae and P. aeruginosa (per lab bench) 2 OF deeps with glucose as carbohydrate (green agar, white cap) Mineral oil (clear viscous liquid, clear cap) 1 nitrate broth (yellowish broth, black cap) Oxidase reagent (tetra-methyl-paraphenylene diamine HCl) Nitrate reagents A (sulfanilic acid) and B (dimethyl-alpha-napthylamine) Procedure – Work in pairs These same tests will be performed on your unknowns as well, but keep results separate in your notebook. Note: the OF tubes and the oxidase test are only used on your gram negative unknown. Week 1 A. OF medium – determines the ability of oxidative and/or fermentative metabolism of glucose. 1. Each pair of students will work with one organism, either K. pneumoniae or P. aeruginosa, while the pair on the opposite side of the bench will work with the other organism. Each pair will inoculate 2 tubes of OF medium (green medium in white cap) with one of the organisms (K. pneumoniae or P. aeruginosa) by stabbing down through the middle of the medium with an inoculating needle. Note: each pair of students have 6 OF tubes for this lab – one set of tubes for their gram- unknown, one set of tubes for their lab partner’s gram- unknown, and the last set of tubes for their assigned organism (K. pneumoniae or P. aeruginosa). 2. Cover one of the tubes with approximately 1 cm of mineral oil (clear capped tube) by pouring the oil aseptically into the tube. Leave the other tube open to the air. 3. Tubes will be incubated at 37°C for 48 h. 4. After incubation, observe the tubes. The OF medium contains bromthymol blue, a pH indicator. It is green at a neutral pH, yellow at an acidic pH, and blue at a basic pH. If the organism can break down the glucose under the given conditions (i.e. in the presence or absence of oxygen), acid is produced and the tube (or a portion of the tube) will change from green to yellow. If acid is produced in both the open and closed (covered with oil) tube, the metabolism of the organism is described as oxidative and fermentative. If acid is produced in only the open tube, the organism is oxidative and the organism only grows by respiration. If acid is only produced in the closed tube, the organism is fermentative and anaerobic. If no growth occurs, the organism cannot utilize glucose or other required nutrients are missing (or the tube was inoculated incorrectly). 5. Record observations and results of both K. pneumoniae and P. aeruginosa. Possible results: 1) organism is oxidative, 2) organism is oxidative & fermentative, or 3) there was little or no growth in either tube. (There are no anaerobes being used in the MB 303 lab, so there are no fermentative only organisms.) B.

Nitrate broth - determines the ability to reduce nitrate to nitrite or nitrogen gas by anaerobic respiration. 1. Each pair of students will inoculate a tube of nitrate broth (black cap tube) with one of the organisms (K. pneumoniae or P. aeruginosa) using an inoculating loop. The opposing pair of students will inoculate the alternate organism into a tube of nitrate broth. 2. Tubes will be incubated at 37°C for 48 h. TAs will incubate uninoculated control tubes as well. 3. After incubation, agitate each tube of inoculated nitrate broth* and then add 4 drops of nitrate A reagent (sulfanilic acid), immediately followed by 4 drops of nitrate B reagent (dimethyl-alpha-napthylamine). In the presence of nitrite, these reagents cause the culture to turn red, indicating that nitrate reduction has occurred. If nitrate has been reduced to nitrite during anaerobic respiration, a red color is seen within 20 minutes Some reactions occur instantaneously and dissipate quickly. Closely observe your tube for the first few minutes after adding reagents A and B. If there is no reaction then check the tube

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4.

5.

again after 20 minutes. Note: a red or rust colored flocculation that was not present before the addition of reagents A+B counts as a positive reaction. If the medium does not turn red after 20 minutes, add a small amount of zinc dust. Zinc causes nitrate to be converted to nitrite, so a red color will result if the nitrate is still present in the medium because the organism could not reduce it. The color will typically develop within a few minutes. If no color change occurs after the addition of zinc, the bacteria reduced nitrate all the way to nitrogen gas. See Figure 14.1 for overview of possible reactions. Record observations and results of both organisms plus the 2 uninoculated control tubes set up by the TAs (1 tube has reagents A+B, 1 tube has A+B plus zinc dust). Possible results: 1) reduction of NO3 to NO2, 2) reduction of NO3 to N2, or 3) no reduction of nitrate. *Note: cell pigmentation can interfere with interpretation of the nitrate test. Be sure to note the presence of any red or pink color in the tube before adding reagents.

Figure 14.1: Overview of Possible Nitrate Broth Reactions

Nitrate reagents A + B Broth

red color forms (NO3 reduced to NO2)

no red color forms

zinc

red color forms (NO3 not reduced) no red color forms (NO3 reduced to N2)

Week 2 C. Oxidase test – determines the presence of the cytochrome c oxidase enzyme, using isolated colonies on a TSA plate or the filter paper method. 1. TSA plates of K. pneumoniae and P. aeruginosa will be provided for oxidase testing. The test works best on plates no older than 24 h, so we have provided freshly streaked plates. 2. Place 1 drop of oxidase reagent (tetra-methyl-paraphenylene diamine HCl) on an isolated colony. Oxidase positive colonies typically exhibit a color change from pink to maroon to black within 5 minutes. Oxidase negative colonies do NOT CHANGE COLOR. A filter paper strip test will be used on unknown cultures that naturally produce red or orange pigmentation. Two spots of culture are placed on the filter paper and then a drop of oxidase reagent is added to one spot. The other spot is left as a negative control, for comparison. A color change to pink/magenta is a positive reaction, indicating the presence of the cytochrome c oxidase enzyme. No color change indicates absence of the enzyme. 3. Record observations and results of both K. pneumoniae and P. aeruginosa. Lab Notebook  Observe and record results of all tests and all organisms (not just the ones that you inoculated) in a neatly labeled table (you may use the table on the next page and staple it into your lab notebook, or design your own in your lab notebook). Be sure to indicate test, organism, observations, and interpretation. Do not use + and – for observation or interpretation! An observation is what you observe and needs to be complete. If it involves a color change then the initial color should be indicated, as well as the final color. If reagents needed to be added, this should also be indicated. An interpretation explains the observed observation and also needs to be complete, explaining the ability (or lack of ability) of the organism (i.e. “the organism has the enzyme X” or “the organism lacks the ability to ferment sugar Y”). 5 pts.  Have your TA initial your completed table before leaving lab. You may re-do your table in a typewritten fashion if you wish, but full credit will only be awarded for your original handwritten observations & interpretations as verified by your TA. 3 pts. Note: All unknown results should be recorded in a separate table in an Ex. 13. results section.

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Results Table for Exercise 14 Klebsiella pneumoniae Observations

Pseudomonas aeruginosa

Interpretation

Observations

OF Medium

Nitrate Broth

Oxidase

Uninoculated Control Tubes Reagents A & B added

Reagents A & B, plus zinc added

Nitrate Broth

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TA initial:

Interpretation

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EXERCISE 15. ADDITIONAL TESTS FOR GRAM POSITIVE BACTERIA Introduction Organisms that live in the presence of oxygen must contain enzymes to protect them from toxic byproducts of oxygen metabolism such as hydrogen peroxide and superoxide. One such enzyme is catalase, which breaks down hydrogen peroxide to water and oxygen. The catalase test is used to separate the staphylococci from the streptococci, enterococci, and lactococci. Streptococci, enterococci, and lactococci are aerotolerant anaerobes that use an alternate enzyme, peroxidase, to break down H2O2. Since no oxygen is evolved from the breakdown of H2O2 by peroxidase the test allows for easy identification of the bacteria that possess catalase. H2O2

peroxidase

2H2O

NADH + H+

H2O2

NAD catalase

H2O + ½O2

Only small molecules cross the cell membrane. In order for bacteria to utilize larger molecules, they must breakdown these molecules outside the cell with the aid of exoenzymes. Exoenzymes are enzymes that are released to the outside of the cell and are commonly used by gram positive bacteria. The Methyl Red (MR) test, applied in combination with the Voges-Proskauer (VP) test, is used to identify bacteria on the basis of their end products formed from metabolism of glucose. Most bacteria metabolize glucose to pyruvate, which is subsequently broken down to a variety of acids (the mixed acid pathway) or to butanediol (the butylene glycol pathway). Methyl red is a dye indicator that changes color at a pH below 4.5, which occurs when bacteria produce acid end products from fermentation. The Voges-Proskauer (VP) test detects the fermentation intermediate acetoin, a neutral compound. MR and VP are tested from the same medium. The tests are commonly used to differentiate Enterics (gram negative bacteria of the intestines) but can be used to separate bacteria from other groups as well, including the gram positive bacteria. Some gram positive bacteria can utilize the disaccharide lactose, particularly any belonging to the Lactic Acid Bacteria group. The ability to ferment lactose will be tested by looking for acid production (a fermentation end product) in tubes of lactose broth that contains only lactose as a carbon and energy source. Bromcresol purple is the indicator. It is purple at neutral pH and yellow at acid pH so the tubes will start as a purple color and become yellow, if lactose fermentation has occurred. Mannitol Salt Agar (MSA) is commonly used to isolate and differentiate the staphylococci and closely related organisms. The medium is selective due to the high (7.5%) salt content, which inhibits gram negative bacteria and many gram positives. Salt tolerant bacteria that can ferment mannitol will produce acids, causing the typically pinkish medium to change to yellow, due to the presence of the pH indicator phenol red. MSA was described in detail in class for Ex. 10 – refer to the notes taken in regards to specific medium ingredients and their roles for more information. Objectives Each student should write appropriate lab objectives in their lab notebook.

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Materials/Organisms per Pair 1 broth culture: Bacillus subtilis or Staphylococcus epidermidis TSA plates inoculated with Enterococccus faecalis and Staphylococcus aureus (per lab bench) 1 MRVP broth (yellowish broth, brown cap) 3% H2O2 Procedure – Work in pairs These same tests will be performed on your unknowns as well, but keep results separate in your notebook. Note: the catalase test is only used on your gram positive unknown. A.

Catalase test – determines the presence of the catalase enzyme. 1. TSA plates of E. faecalis and S. aureus will be provided for catalase testing. The test works best on plates no older than 24 h, so we have provided freshly streaked plates. 2. Place 1 drop of 3% H2O2 reagent on an isolated colony. Immediate bubbling indicates a positive test for the catalase enzyme, due to the enzymatic conversion of hydrogen peroxide to oxygen and water. No bubbling indicates an organism that lacks the catalase enzyme. 3. Record observations and results of both E. faecalis and S. aureus.

B.

MRVP broth (brown capped tube) – determines the types of end-products produced after fermentation of glucose. 1. Each pair of students will inoculate an MRVP broth tube with one of the organisms (B. subtilis or S. epidermidis), using an inoculating loop. The opposing pair of students will inoculate the other organism into an MRVP broth tube. 2. Tubes will be incubated at 37°C for 48 h. 3. After incubation, follow the instructions below for performing the MR and VP tests: Pour ~1 ml of the inoculated MRVP broth into a serological tube for the Voges-Proskauer (VP) test. The ≈9 ml remaining in the original tube will be used for the methyl red (MR) test. (1) Methyl red test – determines the presence of acid end products (i.e. lactic, acetic, formic acid), indicating use of the mixed acid fermentation pathway. Add 10 drops of methyl red to the original tube. Methyl red is a pH indicator that is red at a pH of 4.4 or below and yellow at a pH of 6.2 and above. The development of a red color indicates the presence of mixed acid fermentation products. If acid has not been produced the broth will stay yellow. (2) Voges-Proskauer test – determines the presence of acetoin (acetylmethyl-carbinol), a precursor of 2,3 butanediol, indicating use of the butylene glycol pathway. Add 5 drops of VP I reagent (alpha-napthol) and 5 drops of VP II (40% KOH) reagent to the 1 ml in the serological tube. (If you have more than 1 ml in the tube, double the amount of reagents added). Gently tap the tube and place in a 37°C water bath for 30 minutes. A red color, which typically forms at the top of the broth, indicates a positive test for acetoin, a pH neutral intermediate in the butylene glycol pathway. The end product in the pathway is 2,3 butanediol, which is not easily detected. Acetoin, however, will impart a red color in the presence of alphanapthol and potassium hydroxide, due to its’ oxidation to a diacetyl compound. If acetoin has not been produced the broth will stay yellow. Note: it is possible to get a negative reaction for both the MR and the VP tests, indicating that the organism is not breaking down the glucose or is using a completely different type of metabolism. It is not possible to get a positive for both the MR and the VP tests with the MB 303 unknowns, unless contamination has occurred or an error has been made. 4. Record observations and results of both B. subtilis and S. epidermidis.

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Lab Notebook  Observe and record results of all tests and all organisms (not just the ones that you inoculated) in a neatly labeled table (you may use the tables on the next pages and staple them into your lab notebook, or design your own in your lab notebook). Be sure to indicate test, organism, observations, and interpretation. Do not use + and – for observation or interpretation! An observation is what you observe and needs to be complete. If it involves a color change then the initial color should be indicated, as well as the final color. If reagents needed to be added, this should also be indicated. An interpretation explains the observed observation and also needs to be complete, explaining the ability (or lack of ability) of the organism (i.e. “the organism has the enzyme X” or “the organism lacks the ability to ferment sugar Y”). 4 pts.  Have your TA initial your completed table before leaving lab. You may re-do your table in a typewritten fashion if you wish, but full credit will only be awarded for your original handwritten observations & interpretations as verified by your TA. 3 pts. Note: All unknown results should be recorded in a separate table in an Ex. 13. results section.

65

66

Results Table for Exercise 15 Staphylococcus aureus Observations

Enterococcus faecalis

Interpretation

Observations

Interpretation

Catalase Test

Bacillus subtilis Observations

Staphylococcus epidermidis

Interpretation

MR test

VP test

TA initial:

67

Observations

Interpretation

68

EXERCISE 16. ADDITIONAL TESTS FOR GRAM NEGATIVE (ENTERIC) BACTERIA. Introduction The group of Enteric bacteria (gram negative, non-spore forming rods that are facultative anaerobes associated with the intestinal tract of warm-blooded animals) contain a number of closely related bacteria, many of which are pathogens. These organisms give identical results to the tests performed earlier (oxidase, OF, motility, oxygen requirement, and nitrate reduction), so specific tests must be performed to separate these bacteria. These are some of the more common tests used to separate the various genera, although normally more than 10 tests are used to get definitive separation. A. Ornithine Decarboxylation Some bacteria produce enzymes which can remove CO2 from an amino acid in a decarboxylase reaction. Each decarobxylase enzyme targets a specific amino acid. In MB 303 we will test for decarboxylation of the amino acid ornithine, which requires production of the enzyme ornithine decarboxylase by the organism. Decarboxylases are inducible enzymes, only produced under the following conditions: 1) the organism must have the gene for the decarboxylase, 2) the environment must be acidic, 3) the amino acid specific for the decarboxylase must be present. The media for decarboxylase reactions consists of glucose, yeast extract, peptone, the pH indicator bromcresol purple, and the specific amino acid. Glucose is broken down by many of these organisms with the production of acid. Those organisms capable of producing decarboxylases are induced by these acidic conditions. The decarboxylase will break down the specific amino acid into CO2 and an amine. Amines are very basic so the pH will shift back to neutrality. Thus these enzymes allow these bacteria to continue growing instead of being inhibited by the acid. Interpreting the results of the decarboxylase test is complex. If the organism can utilize the glucose in the medium an acid shift occurs, resulting in the medium changing from purple to yellow in color. When acid is produced, an organism that produces the specific decarboxylase will decarboxylate the amino acid present in the medium and the pH will be shifted to alkaline, causing the medium to return to a purple color. The medium will remain yellow if the organism does not produce the specific decarboxylase. No color change will result if the organism does not grow or does not produce acid from glucose, so be sure to have a control for color comparison. B. Urea utilization Urea is a waste product of protein metabolism in many animals and is excreted in the urine. Some bacteria have an enzyme, urease, which breaks down urea into ammonia and carbon dioxide. The pH of uninoculated urea broth is 6.8. During incubation, bacteria that make urease will breakdown the urea producing ammonia, which causes the pH to rise. At an alkaline pH the urea broth will change from a light orange to a dark pink. C. IMViC IMViC is a mnemonic device which refers to the combination of indole, methyl red, Voges-Proskauer and citrate utilization tests. Indole is produced from the breakdown of tryptophan by the enzyme tryptophanase. Methyl red (MR) is a dye indicator that changes color at a pH below 4.5, which occurs when bacteria produce acid end products from fermentation. The Voges-Proskauer (VP) test is for the fermentation product acetoin. MR and VP are tested from the same medium. Citrate utilization, tested using a citrate slant, indicates that the organism makes a citrate permease enzyme (citrase) that allows it to use citrate as a sole source of carbon and energy. The slant will change from green to blue due to the presence of the indicator bromthymol blue, which turns blue at a basic pH. D. Lactose fermentation Some Enteric bacteria can utilize the disaccharide lactose. The ability to ferment lactose will be tested by looking for acid production (a fermentation end product) in tubes that contain only lactose as a carbon and energy source. Bromcresol purple is the indicator. It is purple at neutral pH and yellow at acid pH. Objectives Each student should write lab appropriate objectives in their lab notebook.

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Materials/Organisms per Student 1 broth culture: E. coli, Proteus vulgaris, Enterobacter aerogenes, Klebsiella pneumoniae 1 ornithine decarboxylation deep (purple agar, purple cap) 1 urea broth tube (orangish broth, red cap) 1 tryptone broth tube (clear broth, turquoise cap) 1 MRVP broth tube (yellowish broth, brown cap) 1 citrate slant tube (green agar, yellow cap) 1 lactose fermentation broth tube (purple broth, green cap) unsterile serological tubes Methyl red reagent (per lab bench) Kovac’s reagent (per lab bench) VP I (alpha-napthol) and VP II (40% KOH) reagents (per lab bench) Procedure – Work in a group of 4 These same tests will be performed on your unknowns as well, but keep results separate in your notebook. A.

Ornithine decarboxylase deep (purple medium with purple cap) – determines the presence of the ornithine decarboxylase enzyme. 1. Each student will inoculate one tube of ornithine decarboxylase medium with their assigned organism. The medium is to be stabbed with an inoculation needle down through the middle of the medium to the bottom of the tube. 2. Tubes will be incubated at 37°C for 48 h. 3. After incubation, observe the tubes. Determine if growth is present by holding the tube up to the light and observing for turbidity along the stab line. If the tube is yellow or yellowwhite, in part or in whole, acid products have been produced due to glucose fermentation but ornithine has not been decarboxylated and the organism lacks the ornithine decarboxylase enzyme. If the tube is purple or purple-gray then the ornithine has been decarboxylated and the ornithine decarboxylase enzyme is produced by the organism. Some bacteria will not ferment glucose and the medium will be the same dark purple as the control. Use an uninoculated tube of decarboxylase medium for comparison as a control. 4. Record observations and results for all four known bacteria: E. coli, Proteus vulgaris, Enterobacter aerogenes, and Klebsiella pneumonia. Possible results: 1) glucose only is fermented, but organism lacks the ornithine decarboxylase enzyme, 2) glucose is fermented and ornithine is decarboxylated by ornithine decarboxylase enzyme, 3) no glucose fermentation (growth off other nutrients in medium), or 4) no growth.

B.

Urea broth (red capped tube) – determine the presence of the urease enzyme. 1. Each student will inoculate one urea broth tube with their assigned organism, using an inoculating loop. 2. Tubes will be incubated at 37°C for 48 h. 3. After incubation, observe the tubes. If the medium is dark pink then the urease enzyme is present, causing the breakdown of urea and the resulting production of ammonium, an alkaline product. If the tube is orange or yellow then the urease enzyme is not produced, since the pH is neutral or acidic. The phenol red in the medium is orange at a neutral pH, yellow at an acidic pH, and hot pink at an alkaline pH. 4. Record observations and results for all 4 known bacteria: E. coli, Proteus vulgaris, Enterobacter aerogenes, and Klebsiella pneumonia.

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C.

IMViC – a combination of several different tests (indole, MR, VP, citrate) commonly used to separate the Enteric bacteria. 1. Each student will inoculate the following media with their assigned organism, using a loop: tryptone broth (turquoise capped tube), MRVP broth (brown capped tubes) and citrate slant (green medium with yellow cap). 2. All the tubes will be incubated at 37°C for 48 h. 3. Record observations and results for all 4 known bacteria: E. coli, Proteus vulgaris, Enterobacter aerogenes, and Klebsiella pneumonia, using the following guidelines: a. Indole test of tryptone broth (turquoise capped tube) - determines the presence of the tryptophanase enzyme. Shake the inoculated broth gently. Add 5 drops of Kovac's reagent. A cherry red ring on the top layer of the tube is indicative of the presence of indole, which is produced from the breakdown of the amino acid tryptophan by the enzyme tryptophanase. If indole is not produced then a yellow ring will form on the top layer of the tube, indicating that the organism does not make the enzyme tryptophanase. Note: interpretation can be difficult for pigmented organisms. Be sure to note the presence of any coloration before the addition of Kovac’s reagent and only note the color of the ring on the top layer of the tube, once reagent has been added. If the color of the liquid below is distracting then use a piece of paper to block out its’ appearance. b. MRVP broth (brown capped tube) – determines the types of end-products produced after fermentation of glucose. Pour ~1 ml of the inoculated MRVP culture tube into a serological tube so that you have a separate sample for the Voges-Proskauer (VP) test. There will be ≈9 ml remaining in the original tube. This will be used for the methyl red (MR) test. (1) Methyl red test – determines the presence of acid end products (i.e. lactic, acetic, formic acid), indicating use of the mixed acid fermentation pathway. Add 10 drops of methyl red to the original tube. Methyl red is a pH indicator that is red at a pH of 4.4 or below and yellow at a pH of 6.2 and above. The development of a red color indicates the presence of mixed acid fermentation products, after pyruvate reduction. If acid has not been produced the broth will stay yellow. (2) Voges-Proskauer test – determines the presence of acetoin (acetylmethylcarbinol), a precursor of 2,3 butanediol, indicating use of the butylene glycol pathway. Add 5 drops of VP I reagent (alpha-napthol) and 5 drops of VP II (40% KOH) reagent to the 1 ml in the serological tube. (If you have more than 1 ml in the tube, double the amount of reagents added). Gently tap the tube and place in a 37°C water bath for 30 minutes. A red color, which typically forms at the top of the broth, indicates a positive test for acetoin, a pH neutral intermediate in the butylene glycol pathway. The end product in the pathway is 2,3 butanediol, which is not easily detected. Acetoin, however, will impart a red color in the presence of alpha-napthol and potassium hydroxide, due to its’ oxidation to a diacetyl compound. If acetoin has not been produced the broth will stay yellow. Note: it is possible to get a negative reaction for both the MR and the VP tests, indicating that the organism is not breaking down the glucose or is using a completely different type of metabolism. It is not possible to get a positive for both the MR and the VP tests with the MB 303 unknowns, unless contamination has occurred or an interpretation error has been made.

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c. Citrate slant (green slant with yellow cap) – determines the presence of the citrate permease enzyme (citrase). Observe slant. A color change of the medium from green to blue denotes a positive for the utilization of citrate as a sole carbon source. A positive test indicates that the organism produces a citrate permease enzyme (citrase) to transport sodium citrate into the cell. Once inside the cell the citrate is utilized, and the excess Na produces a basic change in the medium (blue color). If the slant stays green (no color change) it indicates the absence of the citrate permease enzyme. D.

Lactose broth (green cap with purple broth) – determines the ability of an organism to ferment the carbohydrate lactose. 1. Each student will inoculate one lactose broth tube with their assigned organism, using a loop. 2. Tubes will be incubated at 37°C for 48 h. 3. Observe broths. The broths contain bromcresol purple, a pH indicator that is purple at a neutral pH and yellow at an acidic pH. The presence of a yellow color indicates that lactose was fermented to acid end products, causing a pH change. If lactose is not fermented the tube will remain purple in color. Note: pigments can make interpretation difficult. Be sure to factor in the presence of any pigments known to be produced by the organism.

Lab Notebook  Observe and record results of all tests and all organisms (not just the ones that you inoculated) in a neatly labeled table (you may use the tables on the next 2 pages and staple them into your lab notebook, or design your own in your lab notebook). Be sure to indicate test, organism, observations, and interpretation. Do not use + and – for observation or interpretation! An observation is what you observe and needs to be complete. If it involves a color change then the initial color should be indicated, as well as the final color. If reagents needed to be added, this should also be indicated. An interpretation explains the observed observation and also needs to be complete, explaining the ability (or lack of ability) of the organism (i.e. “the organism has the enzyme X” or “the organism lacks the ability to ferment sugar Y”). 8 pts.  Have your TA initial your completed table before leaving lab. You may re-do your table in a typewritten fashion if you wish, but full credit will only be awarded for your original handwritten observations & interpretations as verified by your TA. 3 pts. Note: All unknown results should be recorded in a separate table in an Ex. 13. results section.

72

Results Table for Exercise 16 Escherichia coli Observations

Proteus vulgaris

Interpretation

Ornithine Decarbox.

Urea Broth

Tryptone Broth

MR test

VP test

Citrate Slant

Lactose Broth

TA initial:

73

Observations

Interpretation

74

Results Table for Exercise 16 Enterobacter aerogenes Observations

Klebsiella pneumoniae

Interpretation

Ornithine Decarbox.

Urea Broth

Tryptone Broth

MR test

VP test

Citrate Slant

Lactose Broth

TA initial:

75

Observations

Interpretation

76

APPENDIX A BACTERIA USED IN MB 303: Alcaligenes faecalis Bacillus cereus B. megaterium B. subtilis Enterobacter aerogenes Entercoccus faecalis Escherichia coli Klebsiella pneumoniae Micrococcus luteus Proteus vulgaris Pseudomonas aureofaciens P. aeruginosa Serratia marcescens Spirillium Staphylococcus aureus S. epidermidis

A-1

APPENDIX B MEDIA USED IN MB 303: Brain Heart Infusion (BHI) Agar Calf brain infusion Beef heart infusion Protease peptone Dextrose NaCl Na2PO4 Agar Distilled H2O pH 7.4

200 g 250 g 10 g 2g 5g 2.5 g 15 g 1000 ml

Intended Use: A rich all-purpose medium for the isolation and cultivation of chemoorganoheterotrophs. Can be used for cultivating fastidious microorganisms, including bacteria, yeast, and molds. Brain Heart Infusion (BHI) Broth Calf brain infusion Beef heart infusion Protease peptone Dextrose NaCl Na2PO4 Distilled H2O pH 7.4

200 g 250 g 10 g 2g 5g 2.5 g 1000 ml

Intended Use: A rich all-purpose medium for the cultivation of chemoorganoheterotrophs. Can be used for cultivating fastidious microorganisms, including bacteria, yeast, and molds. Citrate Agar (aka Simmon’s Citrate Agar) MgSO4 (NH4)H2PO4 K2HPO4 Sodium citrate NaCl Agar Bromthymol blue Distilled H2O pH 6.9

0.2 g 1g 1g 2g 5g 15 g 0.08 g 1000 ml

Intended Use: For the differentiation of gram negative bacteria on the basis of citrate utilization.

B-1

Decarboxylase Medium, L-ornithine Peptone Yeast extract Glucose Bromcresol purple L-ornithine Distilled H2O pH 7.5

5g 3g 1g 0.02 g 5g 1000 ml

Intended Use: For the differentiation of bacteria based on their ability to decarboxylate the amino acid Lornithine, using the enzyme ornithine decarboxylase. Other amino acids can be substituted to test for additional decarboxylases. Lactose Broth Lactose Pancreatic digest of gelatin Beef extract Bromcresol purple Distilled H2O pH 7.0

5g 5g 3g 0.02 g 1000 ml

Intended Use: For the differentiation of bacteria based on lactose fermentation. Lauryl Sulfate Tryptose (LST) Broth Tryptose 20 g Lactose 5g K2HPO4 2.75 g KH2PO4 2.75 g NaCl 5g Sodium lauryl sulfate 0.1 g Bromcresol purple 0.02 g Distilled water 1000 ml pH 6.8 An inverted Durham tube is added to each tube of medium before autoclaving. Intended Use: for the detection of gram negative coliform bacteria in a variety of specimens, based on fermentation of lactose with the resulting production of acid and gas. Mannitol Salt Agar (MSA) Beef extract Proteose peptone NaCl d-Mannitol Agar Phenol red Distilled H2O pH 7.4

1g 10 g 75 g 10 g 15 g 0.025 g 1000 ml

Intended Use: For the isolation and differentiation of staphylococci & closely related organisms based on mannitol fermentation.

B-2

Methyl Red-Voges-Proskauer (MRVP) Broth Buffered peptone Glucose K2HPO4 Distilled H2O pH 7.0

7g 5g 5g 1000 ml

Intended Use: For the differentiation of bacteria based on acid production (methyl red test) or acetoin production (Voges-Proskauer reaction) from the fermentation of glucose. Necessary reagents: methyl red (MR test), -napthol and 40% KOH (VP test). Minimal Salts Medium Na2HPO4 KH2PO4 NaCl NH4Cl Distilled H2O Add after autoclaving: 1 M MgSO4 0.1 M CaCl2 50% glucose pH 7.0

6g 3g 0.5 g 1g 1000 ml 1 ml 1 ml 10 ml

If MSM plates are desired, add 15 g of agar to the broth mixture before autoclaving. Intended Use: For the cultivation of chemoorganoheterotrophs that are not nutritionally fastidious. Motility Medium Heart infusion broth Gelatin Agar Distilled H2O pH 7.2

25 g 53.4 g 3g 1000 ml

Intended Use: For demonstrating motility of microorganisms and for separating organisms in their motile phase. This medium can be used to determine oxygen requirements of an organism as well. Nitrate Broth Beef extract Peptone KNO3 Distilled H2O pH 7.0

3g 5g 1g 1000 ml

Intended Use: For the differentiation of bacteria based on their ability to reduce nitrate. Necessary reagents: sulfanilic acid, α-naphthylamine, zinc powder.

B-3

Ornithine Decarboxylase – see Decarboxylase Medium, L-ornithine Oxidative-Fermentative (OF) Test Medium Tryptone NaCl K2HPO4 Bromthymol blue Agar Glucose Distilled H2O

2g 5g 0.3 g 0.08 g 2g 1g 1000 ml

Intended Use: For the differentiation of a variety of microbes based on their ability to ferment glucose. Peptone Broth Peptone Distilled H2O pH 7.2

5g 1000 ml

Intended Use: For use as a diluent to prevent lysis of cells due to osmotic changes. Plate Count Agar (PCA) Tryptone Yeast extract Glucose Agar Distilled H2O pH 7.0

5g 2.5 g 1g 15 g 1000 ml

Intended Use: For the cultivation, isolation, and enumeration of non-fastidious chemoorganoheterotrophs. RODAC Plates Plate count agar (PCA) + 1% extra agar Intended Use: For the cultivation, isolation, and enumeration of non-fastidious chemoorganoheterotrophs from environmental surfaces. For best results the surfaces must be flat and impervious. Simmon’s Citrate Agar – see Citrate Agar Tryptone Broth Tryptone Distilled H2O pH

10 g 1000 ml

Intended Use: For the differentiation of organisms, especially the Enterobacteriaceae, on the basis of tryptophanase production. Necessary reagents: Kovac’s reagent.

B-4

Trypticase Soy Agar (TSA) Pancreatic digest of casein Papaic digest of soybean meal NaCl Agar Distilled H2O pH 7.3

15 g 5g 5g 15 g 1000 ml

Intended Use: For the cultivation, isolation, and maintenance of a wide variety of fastidious and nonfastidious chemoorganoheterotrophs. Urea Broth Urea KH2PO4 K2HPO4 Yeast extract Phenol red Distilled H2O pH 6.8

20 g 9.1 g 9.5 g 0.1 g 0.01 g 1000 ml

Intended Use: For the differentiation of organisms, especially the Enterobacteriaceae, on the basis of urease production.

B-5

APPENDIX C GLOSSARY OF MEDIA INGREDIENTS Agar: Dried mucilaginous substance extracted from seaweed and used to solidify media. Because of its complex structure, it is difficult for most bacteria to degrade. It melts at 90-100°C and after melting will not re-solidify until 45°C. Beef Extract: An extract of meat used as a nutrient in media, containing amino acids, low molecular weight peptides, nucleotides, organic acids, vitamins, minerals and trace elements. It also contains water soluble proteins and glycogen. This provides energy, all essential macro and micronutrients and growth factors for chemoorganoheterotrophs. Brom Cresol Purple: An indicator of pH range 5.2-6.8. It is yellow at an acidic pH, and purple at a neutral and basic pH. Brom Thymol Blue: An indicator of pH range 6.0-7.6, yellow-green-blue. It is green at a neutral pH, yellow at an acidic pH, and blue at a basic pH. Typically inhibits the growth of gram positive bacteria. Citrate: An intermediate of the TCA cycle. Can be used as a carbon source by some microbes. Mannitol: A sugar alcohol, fermentable by some microbes to yield acid by-products. Peptone: An ingredient used in many complex media, a hydrolyzed protein that contains many amino acids and minerals. The hydrolyzed protein is prepared from partial proteolytic digestion of meat, casein, soyameal, gelatin or other protein source. Serve as sources of energy, C, N, S, and some micronutrients and amino acid growth factors for chemoorganoheterotrophs. Phenol Red: An indicator of pH range 6.6-8.4. It is pink at a neutral pH, yellow at an acidic pH, and hot pink at a basic pH. Proteose Peptone: One of the kinds of peptone. Sodium Lauryl Sulfate: a detergent that inhibits the growth of gram positive bacteria. Tryptone: A pancreatic digest of casein. It contains 12% total nitrogen and 5% NaCl. See peptone. Tryptose: A peptone derived from meat protein. It contains 12% total nitrogen and 6% NaCl. See peptone. Urea: An organic product of protein metabolism. Can be broken down by the urease enzyme to yield ammonia and carbamate. Yeast Extract: An extract of yeast rich in B vitamins and amino acids. It contains numerous unidentified components. Typically it’s used to provide growth factors in a medium.

C-1

APPENDIX D DIFFERENTIAL STAINS A. Capsule stain A capsule is a thick polysaccharide layer located outside the cell wall of some bacteria. Capsules are difficult to stain and are usually observed with a negative stain, where the background is stained with an acidic stain (i.e. congo red) while the cell is stained with a basic stain (i.e. fuchsin). The acidic stain colors the background because the dye is repulsed by the negative charges on the cell wall, while the basic stain colors the cell because it’s attracted to the negative charge on the cell wall. The capsule remains unstained and is seen as a bright translucent area around the cell. Procedure, using congo red and fuchsin: Place a drop of congo red on a slide and mix in a loopful of bacteria. Spread this over the slide so a thin film results. Air dry and warm on hot plate. Do not heat fix. Counterstain with fuchsin. Rinse with water and observe under the microscope. The cells will be pink/red, the background B. Endospore stain Endospores are heat and chemical resistant structures produced by some gram positive bacteria (most notably the genera Bacillus and Clostridium) as a means of surviving adverse conditions. Because endospores are chemically resistant, they are difficult to stain. Heat is used to expand the endospore coat, allowing stain (i.e. fuchsin) to enter the endospore. The cells are then cooled, trapping the fuchsin inside. Cells are rinsed to remove the fuchsin from the vegetative cells, which are then counterstained with an alternatively colored dye (i.e. methylene blue). In this example the endospores are stained pink and the vegetative cells are stained blue. Procedure, using fuchsin and methylene blue: Heat fix a smear of bacteria. Cover smear with fuchsin and steam heat for 5 minutes. Cool for 1 minute. Rinse with water thoroughly. Counterstain with methylene blue. Rinse with water and observe under the microscope. C. Acid-Fast stain (Ziehl-Neelson technique) The acid fast stain is used to differentiate bacteria that contain a high concentration of mycolic acid in their cell wall, such as the Mycobacterium species. Such organisms do not stain well with typical dyes, including the Gram stain dyes. More concentrated dyes such as carbol-fuchsin are used in conjunction with heat over a prolonged period, driving the stain past the cell wall. The bacteria will then be resistant to decolorizing, giving them the designation “acid-fast.” Acid-fast bacteria will be stained red/pink due to retention of the carbol-fuchsin, while any non-acid-fast bacteria will appear blue due to the methylene blue counterstain. Procedure, using carbol-fuchsin and methylene blue: Flood the bacterial smear with carbol-fuchsin dye, heat for 5 minutes over a boiling water bath (adding more dye as needed). Decolorize with acetone-alcohol. Rinse with water. Counterstain with methylene blue. Rinse with water and observe under the microscope.

D-1

APPENDIX F: Example Results Charts for Unknowns (must be typewritten, for full credit!) Jane Doe Unknown #28 Gram positive unknown Test/Medium A. Gram stain B. BHI slant, 37oC

Observation Color & shape (cell arrangement, if it can be determined) Amount and appearance (i.e. color, texture) of growth

C. BHI slant, 30oC

Amount and appearance (i.e. color, texture) of growth

D. Motility

Whether or not growth moved away from stab Location of growth in/on medium Color after reagents A& B, color after zinc (if added) 5 colony characteristics: size, pigmentation, form, margin elevation Presence/absence of bubbles after H2O2 added Growth (if present) & agar color before and after incubation

E. Oxygen requirement F. Nitrate broth G. TSA streak plate

H. Catalase test I.

Mannitol Salts Agar

J. Methyl Red test

Interpretation Gram reaction & morphology Does it grow or not, pigmentation* (indicate if selected temp) Does it grow or not, pigmentation* (indicate if selected temp) Motile or nonmotile Oxygen category Whether or not organism reduces nitrate and if so, to what If culture is pure or not

Presence/absence of catalase enzyme Whether or not organism is salt tolerant. Whether or not organism can ferment mannitol (if known). State if mannitol fermentation ability is not known. Whether or not mixed acid fermentation occured Presence/absence of acetoin

Color before and after reagents, indicate MR reagent added K. Voges-Proskauer test Color before and after reagents, indicate VP I and VPII reagents added L. Lactose broth Color of broth before and after Ability to ferment lactose. incubation Unknown identification = Genus species Exceptions: “none” or list any test that doesn’t match characteristics listed

Note: Attach flow chart and highlight the path used on flow chart. If original flowchart indicated that a “redo” was necessary, a new flowchart must be attached. Note: include observations & interpretations for ALL the tests listed on this page, not just the tests used for final identification.

*white, tan, cream, etc. are all forms of pigmentation. Any color that you can see is a pigment!

F-1

Jane Doe Unknown #28 Gram Negative unknown Test/Medium A. Gram stain B. BHI slant, 37oC

Observation Color & shape (cell arrangement, if can be determined) Amount and appearance (i.e. color, texture) of growth

C. BHI slant, 30oC

Amount and appearance (i.e. color, texture) of growth

D. Motility

Whether or not growth moved away from stab Location of growth in/on medium Color after reagents A& B, color after zinc (if added) Color before and after incubation for both open & closed tubes 5 colony characteristics: size, pigmentation, form, margin elevation Color after oxidase reagent added

E. Oxygen requirement F. Nitrate broth G. OF deeps H. TSA streak plate

I.

Oxidase test

J. Ornithine decarboxylase

Color before and after incubation

K. Urea broth

Color before and after incubation

L. Tryptone broth

Interpretation Gram reaction & morphology Does it grow or not, pigmentation* (indicate if selected temp) Does it grow or not, pigmentation* (indicate if selected temp) Motile or nonmotile Oxygen category Whether or not organism reduces nitrate and if so, to what Oxidative/fermentative category If culture is pure or not

Presence/absence of cytochrome c oxidase enzyme Presence/absence of orthnithine decarboxylase enzyme Presence/absence of urease enzyme Presence/absence of tryptophanase enzyme Whether or not mixed acid fermentation occured Presence/absence of acetoin

Color of ring on top, indicate Kovacs reagent added M. Methyl Red test Color before and after reagents, indicate MR reagent added N. Voges-Proskauer test Color before and after reagents, indicate VP I and VPII reagents added O. Citrate slant Color of slant before and after Presence/absence of citrate incubation permease enzyme P. Lactose broth Color of broth before and after Ability to ferment lactose. incubation Unknown identification = Genus species Exceptions: “none” or list any test that doesn’t match characteristics listed

Note: Attach flow chart and highlight the path used on flow chart. If original flowchart indicated that a “redo” was necessary, a new flowchart must be attached. Note: include observations & interpretations for ALL the tests listed on this page, not just the tests used for final identification. *white, tan, cream, etc. are all forms of pigmentation. Any color that you can see is a pigment! F-2

Appendix G: Lab Summaries This provides a lab-by-lab guideline of things to remember for each lab. In addition, students are always responsible for meeting the detailed requirements set forth in the Lab Notebook section of each exercise. Week 1  Ex. 1: draw 3 organisms in wet mount; make presumptive ID on 2 organisms  Ex. 2: observe 3 prepared slides; show TA your prepared slide (2.5 pts)  Ex. 3: inoculate tubes (4), observe next week  Ex. 4: take sampling bag with you  At end of lab: o Clean microscope, sign sheet, have TA check o Disinfect lab bench, wash hands *Take online safety quiz on Canvas before Lab 2 (10 pts)* Week 2  Lab Notebook due at beginning of lab for grading of Ex. 1 & 2 (12 pts)  Ex. 3: record observations for control tube and E. coli & M. luteus slants, deeps, broths  Ex. 4: prepare PCA plate; incubate RODAC & PCA plate  Ex. 5: prepare/observe Gram stain of mixed culture. Show TA your Gram stain (2.5 pts)  Ex. 6: prepare/incubate streak plate of mixed culture  Ex. 7: prepare dilution scheme (at home)  Ex. 8: scientific writing assignment (at home) Week 3  Ex. 7 dilution scheme due at beginning of lab (5 pts)  Ex. 8 Scientific Writing assignment due at beginning of lab (10 pts)  Ex. 4: observe PCA & RODAC plates, prepare/observe Gram stain; show to TA (2.5 pts)  Ex. 6: observe streak plate of mixed culture  Ex. 7: prepare dilutions, plate, incubate  Ex. 9: observe Winogradsky column layers/colors & 2 organisms in wet mount on videoscope Week 4  Lab Notebook due at beginning of lab for grading of Ex. 3, 4, 5 (22 pts)  Midterm Exam, Ex. 1-6 & 8 (25 pts)  Ex. 7: record plate counts, determine CFU/ml (record on class data sheet), record CFU/ml values for all the students at your lab bench before leaving lab  Ex. 10: prepare 6 streak plates (4 x medium #1, 1 each of media #2, 3) of assigned organism; inoculate medium #4 (broth). Make sure that all media is correctly labeled & incubated!  Ex. 11: determine optimal wavelength (Part A), measure OD of dilutions (Part B)  In-class points: correct clean up/incubation (2.5 pts) Week 5  Lab Notebook due at beginning of lab for grading of Ex. 6, 7, 9 (33 pts)  Ex. 12 in-lab worksheet due before end of lab (5 pts)  Ex. 10: make sure Ex. 10 table is completely filled out (staple into lab notebook)  Ex. 13: Gram stain unknowns (show to TA – 2.5 in-class points); streak 2 BHI slants for each unknown (place 1 at 30oC, 1 at 37oC)

G-1

Week 6  Ex. 13 unknown flow charts due at beginning of lab (20 pts)  Ex. 12: measure OD every 15 min, plate every 30 min; turn in group data sheet with names/seat #s & ODs before end of lab (2.5 pts)  Ex. 13: o Observe BHI slants, determine optimum temperature o Gram +: inoculate BHI slant, nitrate broth, motility tube, TSA plate o Gram -: inoculate BHI slant, nitrate broth, motility tube, TSA plate, OF tubes  Ex. 14: inoculate nitrate broth & 2 OF tubes with K. pneumoniae or P. aeruginosa Week 7  Ex. 12: count colonies; turn in group data sheet with colony counts before end of lab (2.5 pts)  Ex. 13: o Gram +: observe & interpret BHI slant, nitrate broth, motility/oxygen requirement, TSA plate, catalase test. o Gram +: inoculate BHI slant, MRVP broth, MSA plate, lactose broth o Gram -: observe & interpret BHI slant, nitrate broth, motility/oxygen requirement, TSA plate, oxidase test, OF tubes o Gram -: inoculate BHI slant, ornithine decarboxylase broth, urea broth, tryptone broth, MRVP broth, citrate slant, lactose broth  Ex. 14: observe uninoculated nitrate broth tubes; observe & interpret nitrate broth, OF tubes, oxidase test for K. pneumoniae & P. aeruginosa. Get Ex. 14 table initialed by TA.  Ex. 15: inoculate MRVP broth with B. subtilis or S. epidermidis; observe & interpret catalase test for S. aureus & E. faecalis  Ex. 16: inoculate ornithine decarboxylase deep, urea broth, MRVP broth, tryptone broth, citrate slant, lactose broth with E. coli, P. vulgaris, E. aerogenes, or K. pneumoniae *Ex. 10 table and Ex. 11 graphs are due next week. Staple into lab notebook before handing it in* Week 8  Lab Notebook due at beginning of lab for grading of Ex. 10, 11 (36 pts)  Ex. 13: o Gram +: observe & interpret BHI slant, MRVP broth, MSA plate, lactose broth. Streak TSA plate (5 pts) o Gram -: observe & interpret BHI slant, ornithine decarboxylase deep, urea broth, tryptone broth, MRVP broth, citrate slant, lactose broth. Streak TSA plate (5 pts) o Perform mixed Gram stain of both unknowns, show to TA (5 pts)  Ex. 15: observe & interpret MRVP broth for B. subtilis & S. epidermidis. Get Ex. 15 table initialed by TA.  Ex. 16: observe & interpret ornithine decarboxylase deep, urea broth, MRVP broth, tryptone broth, citrate slant, lactose broth for E. coli, P. vulgaris, E. aerogenes, & K. pneumonia. Get Ex. 16 table initialed by TA. Week 9  Ex. 12 assignment due at beginning of lab (40 pts)  Ex. 13 unknown reports due at beginning of lab (62.5 pts) Week 10  Lab Notebook due at beginning of lab for grading of Ex. 13 (raw data), 14, 15, 16 (47 pts)  Lab Final Exam, cumulative (40 pts)

G-2

Name: __________________________________________

Seat #: ______________

Name + seat # on front of lab notebook (not front page!) Bound notebook (not 3-ring binder), dedicated to MB 303 Table of contents up to date with appropriate page numbers Exercise 1: Use of the Microscope Drawings of 3 wet mount organisms, correctly labeled Presumptive ID of 2 organisms (genus level), scientific nomenclature Exercise 2: Simple Stains: Microbial Morphology Observations of B. subtilis, S. aureus, Spirillium slides, correctly labeled Total for Ex. 1 & 2: Table of contents/page #s up to date Exercise 3: Transfer of Bacteria Number of colonies on PCA plate Observations for control tube Observations of broth, deep, slant for E.coli & M. luteus Exercise 4: Environmental Sampling Colony counts for RODAC & PCA plates Indication if sanitation was acceptable for each sample Gram stain – with detailed labeling (ID, scientific nomenclature) Colony appearance (5 colony characteristics) for Gram stained colony Presumptive ID for Gram stained colony Explanation about whether the organism would cause concern or not Complete reference information in ASM style Exercise 5: Differential Stains Objectives for exercise Gram stain observations for mixed culture (with detailed labeling) Observations of demo stains: capsule, endospore, acid-fast

1 1 2

____ ____ ____

3 2

____ ____

3

_____

2

____

1 1 3

____ ____ ____

2 2 1 1 1 1 1

____ ____ ____ ____ ____ ____ ____

2 1 3

____ ____ ____

_____/12 pts

Total for Ex. 3, 4, 5: Table of contents/page #s up to date Exercise 6: Pure Culture: Streak Plate Objectives Observations for streak plate, with 5 colony characteristics, both organisms Indication if isolation was obtained or give reasons why not Exercise 7: Quantification of Bacteria Objectives Appearance of colonies on Petrifilm™ Raw colony counts (individual) in neatly labeled table Calculation of individual CFU/ml, using appropriate values Complete copy of class data for bench in neatly labeled table Calculation of average CFU/ml using class data Comparison of student CFU/ml to class average CFU/ml Exercise 9: Culturing the Unculturable Objectives Record of layers/colors of specific Winogradsky column Identification of Winogradsky column (location/date) Reaction & organism responsible for 2 colors/layers (w/non lab manual ref) Complete reference information in ASM style Observations of 2 different microbes on video scope, with labeling Total for Ex. 6, 7, 9

_____/22 pts 2

____

2 2 2

____ ____ ____

2 1 2 2 2 2 2

____ ____ ____ ____ ____ ____ ____

2 2 1 4 1 2

____ ____ ____ ____ ____ ____ _____ /33 pts

Table of contents/page #s up to date Exercise 10: Culturing Bacteria Objectives for exercises Complete table (pg. 35) with detailed results for all 6 organisms Has title & legend, indicating identity of all 4 types of media Indicates temperature & time of incubation for each medium Student’s organism is circled Exercise 11: Introduction to the Spectrophotometer Objectives for exercise List of OD measurements of methylene blue at diff wavelengths Computer-made graph of wavelength (x-axis) vs. OD (y-axis) List of OD measurements for methylene blue dilutions Computer-made graph of molarity (x-axis) vs. OD (y-axis) List of molarities for original methylene blue & dilutions Calculations of molarities (original plus dilutions) Explanation of what was learned from Part I graph Explanation of whether molarity graph follows Beer’s Law

2

____

2 6 2 1 1

____ ____ ____ ____ ____

2 2 4 2 4 2.5 2.5 1 2

____ ____ ____ ____ ____ ____ ____ ____ ____

Total for Ex. 10, 11: Table of contents/page #s up to date Exercise 13: Identification of Unknowns Observations/interpretations for all tests, written in lab notebook Observations for Gram stain #1 & #2 with labeling, written in notebook Observations of BHI slants for each week (4 weeks total), written in notebook Exercise 14: OF, Anaerobic Respiration Objectives for exercises Results table initialed by TA in Week 7 Ex. 14 results table completed for OF, nitrate broth, oxidase – must list reagents, before/after media color, enzymes produced, etc. - complete observations for K. pneumonaie & P. aeruginosa - complete interpretations for K. pneumonia & P. aeruginosa Observations of nitrate broth uninoculated control tubes Exercise 15: Additional Gram + Tests Objectives for exercises Results table initialed by TA in Week 8 Ex. 15 results table completed for catalase test, MR, VP – must list reagents, descriptions, enzymes produced, etc. for all 4 organisms (B. subtilis, S. epidermidis, S. aureus, E. faecalis) - complete observations for all 4 organisms - complete interpretatations for all 4 organisms Exercise 16: Additional Gram - Tests Objectives for exercises Results table initialed by TA in Week 8 Ex. 16 results table completed for urea, typtone, MR, VP, citrate, lactose, ornithine – must list reagents, before/after media color, enzymes produced, etc. for all 4 organisms ( E. coli, P. vulgaris, E. aerogenes, K. pneumoniae) - complete observations for all 4 organisms - complete interpretations for all 4 organisms Total for Ex. 14, 15, 16, 17

_____ / 36 pts 2

____

5 4 4

____ ____ ____

2 3

____ ____

2 2 1

____ ____ ____

2 3

____ ____

2 2

____ ____

2 3

____ ____

4 4

____ ____ _____ /47 pts