Taylor's University Introduction to Microbiology Practical Manual PDF 2024

Introduction to Microbiology MIC60104 September Semester 2024 Practical Manual STUDENT NAME : __________________________ STUDENT ID : __________________________ PRACTICAL SESSION : __________________________...

Introduction to Microbiology MIC60104 September Semester 2024 Practical Manual STUDENT NAME : __________________________ STUDENT ID : __________________________ PRACTICAL SESSION : __________________________ 1 Koch’s Postulates- Why pure cultures are important! Bacteria are normally found existing with other bacteria and other organisms that live in the community the bacteria inhabit. The diverse population of bacteria will also be great, not just species, but many different genera as well. To identify a particular species a pure culture of that species must be used. It is particularly important in identification of disease causing bacteria. To begin identification, the microbiologist must culture a pure culture (one that comes from a single bacterial cell) from the variety obtained from whatever media is under scrutiny. Robert Koch, one of the fathers of medical microbiology, was the first to put great emphasis on the pure culture. The procedure defined below that enables a specific pathogenic organism to be identified is known as Koch’s Postulates. Koch’s Postulates: 1. The researcher must isolate a pure culture. 2. The isolated organisms must show up in every case of the disease. 3. The same organisms must be recovered in pure culture from a previously healthy, test animal that was exposed to the said organisms and then contracted the disease. Module coordinator : Dr Lai Zee Wei Email : [email protected] Module lecturer : Dr Jason Lee Khai Wooi Email : [email protected] 2  MIC60104 Introduction to Microbiology (Practical) CONTENT Page Laboratory Program and Assessment Information 4 Practical 1 : Microscopy and staining techniques for microbial observations 11 Practical 2: Microbial culturing techniques 23 Practical 3: Microbial analysis and testing techniques 22 Practical 4: Quantification of microbial concentration 37 Laboratory Worksheets 41 Practical Worksheet Practical 1 x Practical 2 x Practical 3 ✓ Practical 4 ✓ 3 Laboratory Component and Assessment Information General Information The laboratory program for MIC60104: - Introduction to Microbiology is designed to illustrate and reinforce important aspects of basic techniques in handling, culture and observation of microorganisms. Having completed the laboratory program, you should be able to: Work independently and responsibly in the laboratory Work effectively as a member of a small team Demonstrate safe handling of microorganisms Perform laboratory procedures according to written protocols Present experimental data in an appropriate graphical or tabular format Prepare a scientific report Identify microbial structures and be familiar with the compound light microscope Performing aseptic procedure in culturing and handling of microorganisms Basic preliminary identification of microorganism through biochemical tests and the use of media The practical component of MIC60104 occupies approximately 12 hours of class time per semester. Attendance at all laboratory sessions is compulsory. Students who cannot attend a session must present evidence, such as a medical certificate to explain their absence. Students who fail to do this will be allocated a mark of zero for missed sessions. Laboratory assessment Marks for laboratory exercises will be allocated as indicated: Laboratory worksheets 20% Final practical examination 20% Practical worksheet (20%) All worksheets must be submitted within 1 week from the date of completion of a practical session. The practical worksheets will be handed back to you with a mark and comments from your lab tutor. You are responsible for keeping your marked papers. Late submission of worksheet will be subjected to penalty of 2% marks reduction per day. Non- attendance at a practical session will result in zero mark for lab quizzes and worksheet unless a medical certificate is supplied. Final Practical Examination (20%) Students will be answering short answer questions of all practical projects. Pre-lab preparation Before attending each practical class you should: Read the practical notes for the experiment to be performed Read the relevant sections from texts and lecture notes 4  Read the accompanying safety sheets so that you are aware of any specific safety concern Students must bring the following to every laboratory session: o Laboratory coat o Fine marker pen Each class begins with introductory talk which explains important aspects of the experiment to be undertaken. Students who missed the talk are not entitled to personal explanations. Students are always encouraged to consult lab demonstrator or lecturer when having difficulty understanding the practical notes. Post-lab Clean up your working bench. Remove all labels from the glassware and place in container provided. Discard contaminated plasticware waste into Biological Hazard Bin and non-contaminated waste into normal dustbin. Discard all contaminated glass slides, cover slips and into container labeled with “glass” on the bench. Clean and wipe off paraffin oil from light microscopes and place it in the designated cabinet. Wash your hands with disinfectant prior to leaving the lab. Plagiarism What is it? Plagiarism occurs whenever you present someone else's work as your own without acknowledgement, and is defined by Taylor’s University (Policy No. TUC-ACA-SOPP-AINT) as follows: "Plagiarism is the action or practice of taking and using as one’s own the thoughts or writings of another, without acknowledgment. The following practices constitute acts of plagiarism and are a major infringement of the University’s academic values: Examples of plagiarism: - Paper is downloaded from an Internet source. - Paper contains part or all of the writings of another person (including another student), without citation. - Paper contains passages that were cut and pasted from an Internet source, without citation. While students are responsible for knowing how to quote from, paraphrase, and cite sources correctly, the ability to apply that information in all writing situations is an advanced literacy skill acquired over time through repeated practice. When a student has attempted to acknowledge sources but has not done so fully or completely, the lecturer may determine that the issue is misuse of sources or bad writing, rather than plagiarism. Factors that may be relevant to the determination between misuse of sources and plagiarism include prior academic integrity education at Taylor’s and the programme level of the student. Lecturers are responsible for communicating their expectations regarding the use and citation of sources What happens if you plagiarize and get caught? The penalties for plagiarism can be severe. They include course failure and exclusion from the University. The School and the University both have a formal hearing process for dealing with plagiarism incidents. The School and University also use detection software for uncovering plagiarism in your assignments. How to avoid plagiarism in your lab books and reports 5 We encourage students to work together in groups when recording and processing data: good team work is important for good science, and discussions with colleagues will help you to identify and solve problems as well as assist the learning process. However, you MUST acknowledge the names of co-worker(s) on all plots, tables, etc. prepared with their assistance. You should ALWAYS write answers to questions IN YOUR OWN WORDS; where words from other sources (e.g. lab manual) are unavoidable, they should be enclosed in quotation marks and the source acknowledged. You should NOT copy answers directly from another student's work - your ability to improve your communication skills will be diminished if you do not make an effort! When marking your reports or laboratory books, if your tutor finds two or more reports with answers in the same wording then that section will be marked as having been plagiarised for all of the students involved. This will typically result in the loss of ALL marks for that section. To avoid plagiarism, you must give credit whenever you: quote or paraphrase from someone’s actual spoken or written words use another person’s ideas, opinions or theories in an assignment or essay make use of pieces of information, such as statistics, graphs, drawings that are not common knowledge How can you avoid unintentional plagiarism? Use quotation marks around everything that comes directly from a text or article Try to summarize ideas and arguments in your own words – don’t just rearrange a few words here and there Check that you have correctly paraphrased the original ideas Check your summary against the original text Use Turnitin to check for copied pieces of text 6  LABORATORY SAFETY RULES The following laboratory safety rules apply specifically for the practical and other activities in the undergraduate teaching laboratories. Ensure that you have completed the Safety Declaration and read the associated Occupational Health and Safety in the Laboratory (Undergraduate Student Edition) Safety Guideline before commencing practical work. For all practical work to be undertaken in this course: Health, safety and environmental aspects of the practical have been considered and risk assessments have been carried out on the chemicals and procedures involved. Safety equipment has been provided where necessary. Appropriate information and supervision necessary for the safe execution of each practical is provided. Specific information about particular hazards and how to avoid, eliminate or minimize your exposure to them is given in the relevant places in the laboratory manual. Students are required to: Avoid, eliminate or minimize hazards of which they are aware. Comply with all occupational health and safety instructions. Make proper use of all safety devices and personal protective equipment. Not knowingly place at risk the health and safety of themselves or any other person. Seek information or advice where necessary or when in doubt before carrying out new or unfamiliar work. Wear protective clothing and footwear. Be familiar with emergency and evacuation procedures. Report and record all accidents and near miss incidents. Emergency Evacuation and Safety Equipment In an emergency and during practice evacuations, move quickly and carefully from the laboratory to the external stairwell or nearest emergency exit. Proceed to the designated assembly area (tutor will advise) and wait there until permission is given to re-enter the building. Never run in the laboratory or along corridors. Be aware of the position of emergency and other exits. Ensure you are aware of the safety facilities of the laboratory including the location and use of safety showers and eyewash stations. Laboratory Dress Code Enclosed footwear must be worn at all times in the laboratory. ‘Enclosed’ requires that the heel and upper foot be covered. You will be denied participation in that session unless wearing suitable footwear. If you are changing shoes before or after the practical, this must be done outside the laboratory. A clean laboratory coat must be worn when conducting practical work and must be fastened (all buttons done up). Sleeves must be rolled down. It must be removed when leaving the laboratory for any reason. You must store your laboratory coat in a plastic bag between practical to reduce the risk of transferring contaminants to your books and other articles. Laboratory coats on which chemicals or biological 7  materials have been spilled must not be taken out of the laboratory and must be left with the Preparation Laboratory staff for decontamination. It is not necessary to wear a laboratory coat in the case of tutorials, exams (except practical exams) or poster sessions. Safety glasses must be worn in practical classes when you are instructed that eye protection is required. Prescription glasses alone are not sufficient – safety overglasses must be worn in addition to these. You are required to provide your own safety glasses or overglasses. Contact lenses do not provide any eye protection for hazardous operations and must be worn in conjunction with approved safety eyewear. If you wear contact lenses during practical involving potential exposure to chemical fumes, vapours or splashes, you should remove them at the first sign of any eye redness or irritation. Where gloves are required, they must be worn. They must be removed if you leave the laboratory for any reason. Long hair must be tied back. If a headscarf is worn, the ends must be tucked into the front of the lab coat. Peaked caps are not permitted in practical classes where gas burners are being used (unless worn backwards) as they constitute a fire risk. Cuts and other skin wounds must be covered. Food and Drink Eating and drinking is prohibited in the laboratory. This includes sweets, gum and drinking from water bottles. Any food or drink must remain in your bag on the bag racks – you may not remove it inside the laboratory. This includes drinking from water bottles while standing at the bag racks. The staff and tutors have authority to dispose of any items of food or drinks as deemed necessary. Take off your laboratory coat and gloves, wash your hands and exit the laboratory with your bag if you need to access food and drinks during the practical. General Rules Leave bags, jackets etc on the bag racks near the entrance to the laboratory. Do not block passageways or fire exits. Keep valuables with you. Do not enter the Preparation Room without permission from laboratory staff or tutors. Keep hands, pens and other material that may become contaminated away from your face. Ensure that workbooks, lab manuals etc do not become contaminated. Mobile phones should be turned off and should not be used during the practical. They should be kept in your bag or pocket and not placed on the bench where they could become contaminated. The use of ear-plug/headphone audio devices (IPods, MP3 players etc) is prohibited in the teaching laboratories. Experiments may only be performed under the direct supervision of a tutor, during the scheduled hours of opening of the laboratory. Follow the instructions of the tutor at all times. Unauthorised experimentation is strictly forbidden. Inappropriate conduct and disruptive behaviour may result in denial of further laboratory access. No reagent, specimen or equipment is to be removed from the laboratory without the permission of the tutor or laboratory officer in charge. Be aware of the conditions required for the safe handling of substances involved in the practical course. Information is provided in the laboratory notes. If in doubt, ask your tutor. Sitting on laboratory benches is prohibited. 8  Pipetting by mouth is prohibited. Automatic pipettes or other pipetting aids are provided. Take care to not contaminate them. Report any defective equipment or broken glassware to the tutor. Spills must be reported to the tutor immediately. A laboratory officer or tutor will clean up if the spill is hazardous. If microbiological cultures are spilled, remain seated and ask someone else to report to the tutor so that the contamination is not spread. Contaminated lab coats must be left with the laboratory officer for decontamination. UV transilluminators and other radiation sources must only be used under the direction of the tutor or a lab officer. Correct use of gas burners will be demonstrated. Be aware of the potential danger of unattended burners as the pilot light is often difficult to see. Before leaving the laboratory, turn the gas burner off at the tap. Handle dissecting equipment with care, store scalpel blades covered and secured inside the dissecting kit and always remove blade from handle. Regard all microbiological cultures as potential pathogens. Work practices should aim to minimize the production of aerosols when working on the open bench. In the case of a practical using microorganism, swab the bench with 70% ethanol when you have finished your work. Waste Disposal Wastes must be disposed of strictly in accordance with the charts displayed in the laboratory, unless you are given special instructions in your laboratory notes or by the tutor or laboratory officer. Do not dispose of waste via the sink unless you are authorised to do so. Hazardous chemical waste is to be disposed of into the correct labeled containers which are provided. Sharps are to be disposed of into the yellow plastic sharps bins provided on the benches. Biological waste should be disposed into biological waste container provided on the bench. Leaving the Lab At the end of the practical session and if you leave the laboratory for any reason during the practical, you must REMOVE your laboratory coat and gloves and wash your hands. Hands-free hand washing sinks and skin disinfectant are located at both the ends of the laboratory bench. If you are changing shoes at the end of the practical, this must be done OUTSIDE the laboratory. The wearing of gloves and/or lab coat when using the water cooler in the corridor is prohibited. Exercise care when opening and closing doors on entering and leaving the laboratory. First Aid Report all accidents, injuries and illnesses to the tutor immediately. If required, trained personnel will administer first aid. All accidents, injuries and illnesses must be recorded. Non injury-causing incidents such as spills, electrical shorts etc must also be reported. Eye injuries (chemical/biological splash or mechanical injury) are serious. Treatment requires immediate and prolonged flushing with water (20 minutes minimum) at the eyewash station. Notify tutor immediately. If contact lenses are worn, remove them as soon as possible. Medical advice should always be obtained for an eye injury – MSDS should accompany student to the medical center. 9  Chemical or biological spills on skin - thoroughly wash affected area with copious quantities of water. Notify tutor immediately. Laboratory officer will consult MSDS to determine appropriate first aid. MSDS should be brought along with the student (if necessary) during the medical treatment. Burns including chemical burns – cool burnt area under running water (10 minutes for thermal burns, 20 minutes for chemical burns). Do not apply ice, lotions or creams. Seek medical advice (take MSDS if chemical burn). Sharps injuries – notify the tutor immediately. Wash the wound and encourage bleeding. If you are feeling unwell or dizzy when participating in a practical class, stop immediately, sit down and notify the tutor. Pregnancy and Allergies Students who are pregnant or trying to fall pregnant may be at higher risk from exposure to certain chemicals and hazards. In addition, some students may develop allergies or may be sensitive to particular chemicals. It is important that you contact the course coordinator or person running the practical class if this applies to you.. 10 Practical 1: Microscopy and staining techniques for microbial observations Objectives: 1. To operate and compare different types of microscopes (light, phase contrast, and dark field) for observing microorganisms using various magnification levels, including the oil immersion technique. 2. To apply simple and differential staining methods for analysing the morphology of microbial cells under the microscope. Introduction The light microscope is a vital tool for examining microorganisms, offering detailed insights into their morphology and behaviour. To fully harness its capabilities, it is essential not only to understand the different types of microscopes, such as light, phase contrast, and dark field, but also to master the techniques for proper setup and maintenance. Additionally, effective staining methods enhance the visibility of microbial cells, allowing for clearer analysis. This practical session aims to provide you with the skills to operate various microscopes and apply both simple and differential staining techniques to observe and analyse microorganisms effectively. Figure 1.1: General anatomy of binocular microscope 11 Descriptions of parts of a binocular microscope and their functions: Binocular Tube This is the housing for lens system. Eyepiece or Ocular lenses This comprises the upper unit of the lens systems. It is a movable unit which is housed in the draw or body tube. Different oculars are used to alter magnification (product of ocular and objective) of the microscope. Diopter adjustment ring The binocular tube has a diopter ring on one side to compensate for differences between the right and left eyes. Revolving nosepiece (quadruple) This is the rotating frame at the base of the binocular tube. It contains four threaded sockets which enable different objectives to be brought into alignment with the ocular lenses. Objective Lens This comprises the lower unit of the lens systems. Several objective lenses are attached to the nosepiece. Course Adjustment Knob This is a large, serrated screw which is attached to the upper part of the limb. Its function is to alter the distance between object under examination and objective lens. Fine Adjustment Knob This is a smaller serrated screw which is attached to the limb. Its function is to finely adjust the distance between the object under examination and objective lens. Stage This is a flat plate attached to the lower part of limb and supports the object under examination. Centrally placed in the stage is a hole which allows light to pass to the object from the light source situated at the base of the microscope. Attached to the stage is a spring-loaded finger which enables a microscope slide to be firmly positioned on the stage. The slide is movable by means of the mechanical stage. Sub-stage Condenser This is attached to lower portion of limb beneath the stage. Fundamentally it is a system of lenses which function is to concentrate light on the object. It contains an iris diaphragm. Iris Diaphragm Complex of inter-leaved metal plates mounted in a frame below the sub-stage condenser. This complex of metal plates can be opened and closed by a small lever to regulate the width of light beam entering the condenser. Variable light control Permits adjustment of brightness. 12 Care of microscope: NOTE: Your tutor will be checking to ensure that you are observing the following rules. a) Always use both hands to carry the microscope. Grasp the microscope arm firmly with one hand and lift it carefully. Place your other hand under the base of the microscope for support as you are carrying it. Keep microscope vertical to ensure the ocular lens does not fall out. b) Each time you use your microscope, clean the optical system (ocular lens, objectives, condenser lens) before and after use. Clean the oil immersion lens last so that you do not transfer oil onto the other lenses. Use only Kimwipe paper to clean the lenses. DO NOT USE FACIAL TISSUES. Avoid touching any of the optical system with your fingers. Skin oils can be difficult to remove. c) Never remove any part of the microscope without informing your lab tutor. d) When using oil immersion, always check and double check that it is the oil immersion (x100) lens that you are lowering into the oil. It is clearly identifiable by a black band around its base. The other objectives are not designed to be used in oil and may be damaged if used in this way. If you accidentally lower the wrong lens into oil, clean it immediately with Kimwipe paper. e) When you have completed your work with microscope, swing the lowest power objective into position. This is to prevent the other two longer objective lens from being accidentally lowered into the condenser. Also set the variable light control to minimum to prolong the life span of the halogen bulb. 13 Section 1: Light microscope Materials: per bench Unstained and stained smears of the following microorganisms: Microorganisms Cellular arrangements Types Simple staining Staphylococcus sp. Grape-like clusters. Gram positive Crystal violet/ bacteria methylene blue Streptomyces sp. Filamentous, branched Gram positive Crystal violet/ structures resembling fungi. bacteria methylene blue Bacillus sp. Rod-shaped chains Gram positive Crystal violet/ bacteria methylene blue Escherichia sp Rod-shaped single cell Gram negative Crystal violet/ bacteria methylene blue Vibrio sp Curved rod Gram negative Crystal violet/ motile bacteria methylene blue Saccharomyces sp. Single, oval cells. Budding yeast Crystal violet/ methylene blue Aspergillus sp. Septate hyphae with conidia Filamentous fungi Lactophenol blue form at the conidiophore tips. Procedures: Examine the microbial cells provided at magnifications of 100x, 400x, and 1000x (oil immersion) using a light microscope. a) Read the rules about care of microscope before you start. You should be able to know the various parts of your microscope as shown in Figure 1.1. Ask your tutor if you are unsure. b) To obtain optimal conditions of illumination for the microscope, use the following procedure: 1. Switch on the microscope. 2. Lower the stage by means of the coarse adjustment and then swing the 10x objective lens into place. Remember: The eyepiece provides 10x magnification. 3. Raise the condenser as far as it can go. The plane surface of the condenser should be almost in line with the level of the stage. 4. Place the slide in the spring-loaded specimen holder. NOTE: The culture smear must be on the upper side of the slide. Take care to release the spring-loaded holder carefully, so as not to break or damage the slide. If any fragments of slide fall onto sliding surfaces of the microscope, damage may result. 5. Focus on the specimen using coarse and then fine focus adjustment knobs. 6. Looking through the binocular tube, adjust the distance between the sliding eyepieces until perfect binocular vision is obtained. 7. Close the iris diaphragm completely and then open it until the field is just fully illuminated. This does not mean that the iris diaphragm needs to be fully open. 14  8. If the intensity of light is too great it should be decreased by turning the voltage control dial. Do not lower the condenser. 9. At this point the PRE-FOCUSING LEVER CAN BE LOCKED to avoid changing the coarse focus while switching to other objectives or changing slides. 10. Without altering the focus, bring the x40 lens into position. Focus on the smear using FINE FOCUS ONLY. Adjust the iris diaphragm to give a well illuminated field. 11. Examine the smear using this optimal illumination at 10X and 40X objective lens. c) Examine specimens using oil immersion microscopy at 1000x magnification. 12. To use the 100x objective lens, place a drop of oil on the dried smear. Without changing the focus, swing the 40x lens away so that the 100x objective lens comes into position and immerses in the oil. Provided you have not disturbed the coarse focus, the field should be visible and only require fine focusing. It is important to turn the fine focus slowly and cautiously to avoid breaking the slide or damaging the lens. d) Identify the clarity of the view and the cellular morphology (size, structure, and arrangement) of both unstained and stained smears of microbial specimens. 15 Section 2: Other types of microscopy A. Phase contrast microscopy Material: Unstained wet preparation of Saccharomyces sp. Procedures: per class Functionally similar to light microscopy, phase contrast microscopes detect and enhance small differences in refractive indices, thereby improving the clarity and contrast of the object being examined. They are especially useful for viewing specimens where little natural contrast exists, such as wet preparations of UNSTAINED material. Beams of light pass through the ANNULAR DIAPHRAGM (Figure 1.2) of the phase contrast condenser and are directed towards the specimen. These beams are partially deflected by the different refractive indices of the specimen’s various densities and thicknesses. The light then passes through the PHASE PLATE (Figure 1.2) in the phase contrast objective lens, where the phase differences are converted into changes in amplitude. Areas of uniform refractive index appear bright, as the light waves are not significantly altered. Conversely, areas with different refractive indices cause a decrease in amplitude, which appears as contrast or darkness. a) The microscope set up for phase contrast microscope is similar to that described in Section 1. 1. Ensure that the phase contrast objective lenses (*Typically labeled with a special ring or designation, e.g., Ph 1, Ph 2) are properly installed on the microscope. 2. Align the correct annulus of the condenser with the corresponding phase contrast objective lens. Proper alignment is crucial for achieving the desired contrast and clarity. b) Identify the clarity of the view and the cellular morphology (size, structure, and arrangement) using both the light microscope and the phase contrast microscope. The lab tutor should set up both microscopes side by side for students to observe and compare the differences. Figure 1.2: Annular diaphragm and phase plate of the phase contrast microscope 16 B. Dark field microscopy Material: Unstained wet preparation of Vibrio sp. Procedures: per class Certain bacteria are so thin and transparent that they have low inherent contrast and cannot be clearly resolved using direct preparations, even with phase contrast microscopy. These bacteria are best visualized using dark field microscopy, a technique where light is scattered or refracted off the surface of the specimen, producing a bright image against a dark background. Spiral organisms, such as Treponema pallidum (which causes syphilis), Leptospira interrogans (which causes leptospirosis), and Vibrio cholerae (which causes cholera), can be readily observed using dark field microscopy, but they may not be as clearly visible under a standard light microscope. a) The microscope set up for dark field microscope is similar to that described in Section 1. 1. Ensure that the phase contrast objective lenses (*Typically labeled with a special ring or designation, e.g., Ph 1, Ph 2) are properly installed on the microscope. 2. Microscope set up for dark field microscopy with a wet preparation. Unstained wet preparation of Vibrio sp. b) Identify the clarity of the view and the cellular morphology (size, structure, and arrangement) using both the light microscope and the dark field microscope. The lab tutor should set up both microscopes side by side for students to observe and compare the differences. Section 3: Simple and differential staining A. Fixed smear and wet mount preparations A “smear” of bacteria or yeast is made on a microscope slide, fixed, stained, dried and, without using a coverslip, usually examined using a microscope with oil immersion technique. A thin layer of microbial smear is essential for uniform staining and easy observation for determining the shape and arrangement of cells. Aseptic technique must be applied when taking samples of a culture for making smear. Passing the dried microbial smear through Bunsen flame is called heat fixing. There is a slight shrinkage of cells during this process which helps bacterial cells to adhere to a glass slide. If the slide is overheated, the cells will warp, and the cell’s structure will be indistinguishable. In contrast, a wet mount is prepared by suspending a specimen in a drop of liquid on a slide and covering it with a coverslip. Material: Escherichia coli on nutrient agar 1 plate Escherichia coli in nutrient broth 1 tube Clean microscope glass slide 3 slides Inoculating loop 1 17 Bunsen burner 1 Permanent marker 1 Procedures: per group a) Preparing a fixed smear from bacterial culture from solid media. 1. Obtain a glass slide, use a permanent marker to draw 2 circles on a glass slide. The slide may be turned over so that the markings (circle drawn using marker pen) are at the bottom of the slide. 2. Place 1 drop of tap water in each of the circles on the glass slide. 3. Without scraping or cutting the agar, obtain a small amount of bacteria (a small portion of a single colony) using a sterile inoculating loop and suspend it with the water on the first circle and spread evenly into a thin smear. Flame the loop. 4. Transfer a loop full of suspended cells from the first circle into the second circle, spread evenly to form a thin layer and flame the loop. 5. Dry the smear by placing it close to Bunsen burner flame (or 1 foot above the flame). 6. After the smear is completely dried, pass the slide with cells facing down through the flame of a Bunsen burner several times to heat fix the smear. This adheres the bacteria to the slide and prepares the specimen for staining. b) Preparing fixed smear from liquid culture can be done similarly as described above using liquid culture without the need to resuspending the culture. After that, follow Section 3, steps (a) 4-6. 18 c) Wet mount is prepared by placing one drop of culture suspension on a slide and covering it with a coverslip. B. Staining The chemistry of staining is based on the principle that opposite charges attract and like charges repel. Most bacteria, when placed in an aqueous environment at a neutral pH (around 7), carry a net negative electrical charge. These negatively charged bacterial cells attract positively charged molecules (cations) and repel negatively charged molecules (anions). Each dye used in microbiology is a salt composed of two ions: a positively charged cation and a negatively charged anion. Either ion can serve as the chromophore, the part of the molecule responsible for its colour. In most commonly used stains (basic dyes), the chromophore is associated with the positively charged cation. Basic dyes, such as methylene blue, crystal violet, and safranin, are attracted to the negatively charged bacterial cell surface, making them ideal for simple staining. Conversely, acidic dyes (where the chromophore is the anion) are repelled by the negatively charged cells. These dyes, like nigrosine, stain the background, leaving the cells unstained and colourless, a technique commonly used in negative staining. There are two broad types of staining techniques: i. Simple Stain: A simple stain involves the application of a single staining reagent to highlight the shape and arrangement of cells. Suitable stains for this method include basic dyes, where the colour-bearing ion (chromophore) is the cation, such as methylene blue, crystal violet, and safranin. ii. Differential Stain: A differential stain uses a series of staining reagents to differentiate between various parts of a cell or between different types of microorganisms. For example, Gram staining distinguishes between Gram-positive and Gram-negative bacteria based on differences in their cell wall structure. This method is crucial for bacterial classification and identification. While yeast cells can be stained by the Gram method, the technique does not provide useful information for their identification. 19 Material: *Mixed culture in nutrient broth 1 tube (Escherichia coli : Staphylococcus aureus-1:1) Clean microscope glass slide 3 slides Inoculating loop 1 Bunsen burner 1 Crystal violet Sharing Safranin Sharing Iodine Sharing Mineral oil Sharing 95% ethyl alcohol Sharing Procedures: per group a) Simple staining. 1. Obtain a fixed smear prepared in Section 3A. 2. Flood the smear (not the whole slide) with crystal violet and leave for 1 minute. 3. Wash with running tap water, blot dry with paper towel and view under microscopy with appropriate magnification. 4. For wet mount staining, the stain is introduced by placing it at the edge of the coverslip of a wet mount to allow the stain to diffuse into the specimen through capillary action. b) Gram staining. In 1884, while studying the etiology of respiratory disease in Streptococcus pneumoniae from human lung tissue, Hans Christian Gram discovered a staining technique that differentiated bacteria during autopsy. This discovery revolutionized microbiology, and Gram’s staining method is now performed millions of times daily worldwide. Gram’s technique categorizes bacteria into two broad groups based on their cell wall properties, Gram-positive and Gram-negative. After the Gram staining procedure, Gram-positive bacteria appear purple, while Gram-negative bacteria appear pink. 20 The principle behind Gram's staining method is the cell's ability to retain the crystal violet stain, with the help of iodine as a mordant, when exposed to a decolorizing agent, typically alcohol or acetone. Bacteria that retain the violet or purple colour are classified as Gram-positive, while those that lose the colour and take up the counterstain (safranin) are classified as Gram- negative, appearing pink or red. 5. Perform fixed smear according to Section 3A using a *mixed culture. 6. Flood the smear with crystal violet, leave for 1 minute, and wash with running tap water. 7. Immediately flood the smear with iodine, leave for 1 minute, and wash under running tap water. 8. Hold the slide at about 45° angle where the smear is clearly visible, apply the decolorizer drop by drop to the top edge of the smear for 5-10 seconds or until the color is faded. 9. Immediately wash with tap water and counterstain with safranin for 1 minute. 10. Wash with running tap water, blot dry with paper towel and view under microscopy with appropriate magnification. 21 Expected observation: Figure 1.2: Gram-stained Escherichia coli and Staphylococcus aureus. Gram positive S. aureus stains purple, while Gram negative E. coli stains pink. 22 Practical 2: Microbial culturing techniques Objectives: 1. To aseptically inoculate microbial cultures on solid and liquid media. Introduction Microbial culturing techniques are critical for studying microorganisms, and successful culturing depends on the application of rigorous aseptic techniques. These practices are essential to prevent contamination of cultures, laboratory personnel, and the surrounding environment. Proper inoculation procedures and adherence to safety protocols ensure the accuracy of experimental results and maintain a sterile workspace. To maintain aseptic conditions: 1. Ensure all necessary equipment and materials are within reach before starting and complete procedures as quickly as possible. 2. Minimize exposure of open culture vessels to the air. 3. Perform subculturing and inoculation near a Bunsen burner flame to utilize the upward air currents. 4. Flame the neck of test tubes or bottles immediately after opening to direct air movement outward, keeping the vessel as horizontal as possible. 5. Limit exposure of sterile surfaces, such as Petri dishes, to the air. 6. Avoid touching or contaminating sterile pipettes with non-sterile surfaces, including clothing and work surfaces. 7. Refrain from talking, sneezing, or coughing near cultures and media. Section 1: Aseptic and microbial inoculation techniques. A. Aseptic techniques Material: Inoculation loop, Bunsen burner, Universal bottle, blue cap bottle, conical flask with cotton stopper, nutrient agar plate, molten nutrient agar, petri dish, 100 µL-pipette and tips, glass spreader/hockey stick, 70% ethanol in beaker. Procedures: per class The lab tutor will demonstrate these techniques, after which students will practice them in the laboratory. Mastery of these methods is essential for the subsequent microbial culturing and testing exercises. 1. Flaming an inoculation loop/needle (Figure 2.1). 2. Flaming the neck of bottles (Figure 2.2). 3. 9-streak streaking method (Figure 2.3). 4. Pour plate inoculation (Figure 2.4). 5. Spread plate inoculation (Figure 2.5). 6. Flaming a glass spreader for spread inoculation (Figure 2.6). 7. Lawn culture method (Figure 2.7). 23 Figure 2.1: Flaming the inoculation loop. The procedure involves gradually heating the loop by first positioning the handle end in the pale blue cone of the flame, which is the coolest part. Slowly draw the rest of the loop into the hottest region of the flame until it becomes red hot, ensuring the entire length of the loop is adequately heated. Allow it to cool before use and re-sterilize immediately after use by repeating the process. 24 Figure 2.2: Flaming the neck of a bottle. Begin by loosening the cap and lifting the bottle with your left hand. Use the little finger of your right hand to remove the cap, turning the bottle instead of the cap. Do not set the cap down. Sterilize the neck by passing it forwards and backwards through the Bunsen flame. Replace the cap with your little finger, turning the bottle rather than the cap. Ensure labels are securely placed in a spot that won’t rub off during handling, using marker pens or self-adhesive labels. 25 Figure 2.3: 9-streak streaking method for isolating bacterial colonies. Begin by flaming the inoculating loop and the opening of the universal bottle. Streak a heavy loopful of bacterial culture in a primary inoculum across the corner of the nutrient agar plate (area A). Flame the loop, allow it to cool, and then rotate the Petri dish 90° anticlockwise. Streak from area A in three parallel lines to area B, carrying over a small amount of culture. Flame the loop, cool it, and turn the dish 90° anticlockwise again. Streak from area B to area C in three parallel lines. Finally, flame the loop, cool it, and turn the dish 90° anticlockwise. Streak from area C to the center of the agar (area D). Close the Petri dish and flame the loop. Incubate the plate inverted at 37°C overnight. Figure 2.4: Pour Plate Method. Inoculate 0.1-1 ml of bacterial suspension into a sterile Petri dish. After removing the lid from the molten agar bottle, flame the neck to maintain sterility. Pour the molten agar into the Petri dish, then gently swirl to ensure the inoculum is well mixed and the agar covers the dish evenly. Avoid contact between the agar and the lid, and minimize air bubbles. Allow the agar to solidify, then incubate the plate inverted at 37°C overnight. Photograph the plate the next day to observe and analyse single colonies. 26 Figure 2.5: Spread plate technique using a glass spreader. Begin by aliquoting 50-200 µl of bacterial culture onto the solid nutrient agar in a Petri dish. Close the lid of the Petri dish and discard the used pipette tip. Dip the lower end of a glass spreader into 70% alcohol and flame it with a Bunsen burner, ensuring it points downwards to avoid burns and keeping the alcohol beaker away from the flame (Figure 2.6). Allow the spreader to cool for 30 seconds without placing it on the bench. Spread the inoculum over the agar surface using top-to-bottom or side-to-side motions, covering the entire surface evenly. Close the Petri dish lid, sterilize the spreader, and let the inoculum dry before incubating the plate at 37°C in an inverted position. Photograph the plate the next day to observe single colonies. Figure 2.6: Flaming a glass spreader. Dip the lower end of the glass spreader into 70% alcohol and then pass it through the Bunsen flame, ensuring the spreader is held downward to avoid burns and keeping the alcohol beaker away from the flame to prevent accidents. 27 Figure 2.7: Lawn culture method. Suspend 2 to 3 single colonies of microbial culture in 10 ml PBS and vortex gently with the lid securely attached until a faintly turbid suspension is obtained. Swab the surface of a nutrient agar plate with this inoculum to achieve an even lawn of inoculum. 28 B. Microbial inoculation and growth Material: Inoculation loop, Bunsen burner, Universal bottle, blue cap bottle, conical flask with cotton stopper, nutrient agar plate, molten nutrient agar, petri dish, 100 µL-pipette and tips, glass spreader/hockey stick, 70% ethanol in beaker. Media: Tryptic soy agar (TSA) Eosin Methylene Blue agar (EMB) MacConkey agar (MCA) Hektoen Enteric agar (HE) Sheep Blood Agar (BA) Procedures: per group Microorganisms can be cultured in laboratory settings for various purposes, including general propagation, microbial differentiation, or identification. All-purpose (complex) media, such as Tryptic Soy Agar (TSA), Tryptic Soy Broth (TSB), Nutrient Agar (NA), Nutrient Broth (NB), and Blood Agar (BA), are commonly used to support the growth of a wide variety of microorganisms, including pathogens and normal human flora. In contrast, differential media are extensively used in microbiological labs, especially in food, pharmaceutical, and medical industries, to distinguish between different bacterial genera. Selective media, containing dyes or chemicals that inhibit the growth of certain microorganisms, help in the isolation and differentiation of particular genera by promoting distinct colony colors or growth patterns. 1. By using techniques learned in Section 1A, inoculate the microbial cultures on respective media. 2. Observe colony colour on agar surface after incubation. Media Microorganisms Aseptic techniques Growth condition TSA Escherichia coli Streak plate 37°C, 24 hours EMB Escherichia coli Streak plate 37°C, 24 hours Enterobacter aerogenes Pseudomonas aeruginosa Staphylococcus aureus MCA Escherichia coli Streak plate 37°C, 24 hours Enterobacter aerogenes Pseudomonas aeruginosa Staphylococcus aureus HE Escherichia coli Streak plate 37°C, 24 hours Salmonella Shigella BA Streptococcus pyogenes Streak plate 37°C, 24 hours Streptococcus pneumoniae Staphylococcus epidermidis 29 Expected growth and observations: Eosin Methylene Blue agar (EMB) EMB, the Levine formulation, is commonly used to study enteric bacteria, particularly for distinguishing between lactose-fermenting and non-fermenting microorganisms. This medium is widely applied in medical bacteriology, as well as in methods recommended by the American Public Health Association (APHA) for detecting and enumerating coliforms, which are common contaminants in food and drinking water. Peptone serves as a nutrient source to support bacterial growth, while sucrose and lactose act as fermentable carbohydrates. The inclusion of sucrose allows the detection of coliforms that ferment sucrose more readily than lactose. Eosin and methylene blue dyes serve two functions, acting as partial inhibitors of Gram-positive bacteria while also functioning as pH indicators. The medium is differential because of its ability to distinguish between lactose fermenters and non-fermenters based on color changes in colony appearance. For instance, lactose-negative bacteria such as Pseudomonas aeruginosa, Salmonella and Shigella form translucent, amber, or colorless colonies. In contrast, coliforms that ferment lactose and/or sucrose produce blue-black colonies with dark centers, often exhibiting a characteristic green metallic sheen. Organisms like Enterobacter may form mucoid pink colonies because Enterobacter ferments lactose, but it does not produce the same dark purple or metallic green sheen seen in strong lactose fermenters like Escherichia coli. The pink color indicates lactose fermentation, and the mucoid appearance is due to the production of a capsule. Enterococcus faecalis strains are partially inhibited on EMB, appearing as colorless colonies due to their inability to ferment lactose or sucrose. The methylene blue in EMB selectively inhibits Gram-positive bacteria, ensuring that primarily Gram-negative bacteria grow on this medium. Thus, EMB agar is effective in selecting Gram-negative bacteria through methylene blue inhibition while also distinguishing lactose-fermenting from non-fermenting bacteria based on colony color. Microorganisms Growth Colony color Escherichia coli Good (Strong acid) (+) Green metallic sheen Enterobacter aerogenes Good (Weak acid) (+) Pink Pseudomonas aeruginosa Good (-) Colorless/amber Staphylococcus aureus Inhibited (Gram +ve) No growth MacConkey agar (MCA) MacConkey agar is used to differentiate between lactose-fermenting (coliform) and non-lactose- fermenting (enteric) bacteria. Lactose-fermenting bacteria, often non-pathogenic coliforms, produce pink to red colonies due to acid production, which lowers the pH and interacts with the neutral red dye in the medium. In contrast, non-lactose-fermenting bacteria, which may include pathogenic enteric species, form colorless or slightly opaque colonies because they do not produce acid from lactose fermentation. The addition of bile salts and crystal violet to the agar makes it selective by inhibiting the growth of Gram- positive bacteria, thereby allowing only Gram-negative bacteria to thrive. Microorganisms Growth Colony color Escherichia coli Good (Acid) Pink, non-mucoid Enterobacter aerogenes Good (Strong acid) Pink, mucoid, neutral red turns yellow Pseudomonas aeruginosa Poor (Uses peptone, produces Colorless/opaque ammonia, basic pH) Staphylococcus aureus Inhibited (Gram +ve) No growth 30 HE Agar Hektoen Enteric (HE) agar is a selective and differential medium designed to isolate and differentiate pathogenic enteric bacteria, such as Salmonella and Shigella, from non-pathogenic coliforms and enteric bacteria. It is particularly effective for isolating these pathogens in stool samples. HE Agar contains bile salts, bromothymol blue, and acid fuchsin, which inhibit the growth of most Gram- positive bacteria and select for Gram-negative organisms. Lactose-fermenting bacteria produce orange to pink colonies due to the acidification of the medium. In contrast, bacteria that produce hydrogen sulfide (H₂S) generate black colonies or black precipitates within the agar due to the reaction between thiosulfate and ferric ammonium citrate in the medium. The differentiation of pathogenic enteric bacteria from non-pathogenic ones is achieved through these color changes: lactose fermenters will show orange or pink colonies, while H₂S producers will form black colonies or have black centers, indicating hydrogen sulfide production. This selective and differential medium facilitates the identification of enteric pathogens and helps in distinguishing them from other coliforms. Microorganisms Growth Colony color Escherichia coli Good (Ferments lactose, H2S Orange yellow colonies, may have not produced) bile precipitation. Shigella sp. Good (Not ferment lactose, Green/light green colonies H2S not produced) Salmonella sp. Good (Not ferment lactose, Blue-green with black centers H2S produced) Sheep Blood Agar (BA) Sheep blood agar is a versatile medium widely used for isolating and differentiating bacterial species based on their hemolytic activity, especially among Streptococcus species. This medium supports the growth of a broad range of bacteria and facilitates the observation of their hemolytic properties. This ability to classify bacteria based on their hemolytic behavior is crucial for accurate identification and characterization in clinical and research settings. Hemolytic activity is categorized into three types on blood agar: i. Alpha-hemolysis (Streptococcus pneumoniae) Bacteria causing alpha-hemolysis reduce hemoglobin to methemoglobin, producing a greenish discoloration around the colonies. This indicates partial lysis of red blood cells. ii. Beta-hemolysis (Streptococcus pyogenes) Bacteria that exhibit beta-hemolysis lyse red blood cells and degrade hemoglobin, resulting in a clear, colorless zone surrounding the colonies. This indicates complete lysis of red blood cells. iii. Gamma-hemolysis (Staphylococcus epidermidis) Bacteria that do not produce any change in the appearance of the medium are classified as gamma-hemolytic. There is no discoloration or lysis of red blood cells around these colonies. 31 Practical 3: Microbial analysis and testing techniques Objectives: 1. To assess antibiotic susceptibility using fresh lawn cultures. 2. To classify bacteria using rapid biochemical tests Introduction In this practical, students will explore fundamental testing techniques used in microbiology. The first objective involves assessing the antibiotic susceptibility of bacteria through the use of fresh lawn cultures, a common method to determine bacterial resistance or sensitivity to antibiotics. The second objective focuses on the classification of bacteria using rapid biochemical tests, enabling the identification of specific bacterial species based on their metabolic properties. These techniques are essential for understanding microbial behavior and informing appropriate antimicrobial treatments. Section 1: Antimicrobial susceptibility test: Disc diffusion method Material: Nutrient agar 5 plates Sabouraud dextrose agar 1 plate Sterile swabs 6 pieces Sterile antibiotic disc dispenser 1 unit Phosphate buffered saline (PBS) 10 ml Antibiotic discs: Tetracycline 30 µg Ampicillin 10 µg Erythromycin 15 µg Gentamicin 10 µg Chloramphenicol 30 µg Nystatin 10 µg Procedures: per group Isolating an infectious agent from a patient with a disease is often not sufficient for determining the appropriate therapy, as many organisms exhibit resistance to antimicrobial agents. Since the susceptibility of most bacteria, fungi, and viruses to antimicrobial agents cannot be predicted, testing individual pathogens against specific antimicrobials is often necessary. Several techniques are available for routine testing of antibiotic sensitivity. One of the most commonly used methods is the disc diffusion assay. In this method, filter paper discs containing standard concentrations of various antimicrobial agents are placed onto a freshly inoculated lawn cultures. The antimicrobial agents diffuse radially from the discs, and the zone of inhibition around each disc is measured. This zone indicates the susceptibility or resistance of the bacteria to the antimicrobial agent. 1. Using the techniques learned in Section 1 (Practical 2), prepare lawn cultures for the disc diffusion test according to the cultures listed in the table on the next page. 32 2. Using a sterile disc dispenser, under the guidance of a lab tutor, place discs onto the surface of the freshly lawn cultures. 3. Incubate all plates at 37°C for 24 hours. 4. Measure the diameter of the zone of inhibition (ZOI). Resistance, intermediate susceptibility, or susceptibility can be determined by comparing your results to the ZOI standards. Microorganisms Media Aseptic techniques Antibiotic discs Escherichia coli Nutrient agar Lawn culture method Tetracycline, 30 µg Staphylococcus aureus Nutrient agar Lawn culture method Erythromycin, 15 µg Chloramphenicol, 30 µg Penicillin-resistant Nutrient agar Lawn culture method Ampicillin, 10 µg Staphylococcus aureus Gentamicin, 10 µg Pseudomonas aeruginosa Nutrient agar Lawn culture method Candida albicans Sabouraud Lawn culture method Nystatin, 10 µg dextrose agar Zone of inhibition (ZOI) standard Zone diameter interpretative standards (mm) Test cultures Resistant Intermediate Susceptibility Tetracycline (30 µg) S. aureus ≤14 15-18 ≥19 E. coli ≤14 15-18 ≥19 S. pneumoniae ≤18 19-22 ≥23 P. aeruginosa ≤14 15-18 ≥19 Erythromycin (15 µg) S. aureus ≤13 14-22 ≥23 E. coli - - - S. pneumoniae ≤15 16-20 ≥21 P. aeruginosa - - - Chloramphenicol (30 µg) S. aureus ≤12 13-17 ≥18 E. coli ≤12 13-17 ≥18 S. pneumoniae ≤20 - ≥21 P. aeruginosa - - - Ampicillin (10 µg) S. aureus ≤28 - ≥29 E. coli ≤13 14-16 ≥17 S. pneumoniae - - ≥24 P. aeruginosa - - - Gentamicin (10 µg) S. aureus ≤12 13-14 ≥15 E. coli ≤12 13-14 ≥15 S. pneumoniae - - - P. aeruginosa ≤12 13-14 ≥15 Nystatin (10 µg) C. albicans