Lab-manual-update (1).docx

Laboratory Manual Course No.: CFPE 3212 Course Title: Food Microbiology & Molecular Cell Biology Sessional

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Contents

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Introduction to Microbiology Laboratory &Rules

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Preparation and Sterilization of Simple Culture Media

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Application of Aseptic Inoculation Techniques in Solid and Liquid Culture Media

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Morphological Examination of Microorganisms

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Gram Staining Method: Identification of Gram Positive and Gram Negative Microorganisms with Physiological Examination

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Total Bacterial Count (TBC) of food samples

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Use of beneficial microorganism in producing fermented food product (Dahi)

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Exp. 1: Introduction to Microbiology Laboratory & Rules Laboratory Rules Regard all cultures as potentially dangerous and treat them accordingly. Some of the microorganisms you will be studying in class can cause disease. Therefore, great care should be taken in handling cultures, slides and other materials that have been in contact with living microorganisms. The following precautions should be taken: 1. Only necessary articles should be brought into the laboratory – coats, cases, etc. should not be brought in. 2. Laboratory coats must be worn in the laboratory and MUST be returned to the hangers after use. They should never be worn anywhere other than microbiology laboratories and especially not in food processing area. 3. Safety spectacles or prescription spectacles MUST be worn at all times. 4. DO NOT eat, drink or smoke in the laboratory and never place pipettes, pencils, pens, labels or other materials in your mouth! 5. Sterilise inoculating wires and loops BEFORE and AFTER use by heating in a flame until red-hot. Avoid spattering material by gradually introducing the loop into the flame. 6. DISCARD used plastic/glass disposable pipettes, Pasteur pipettes, microscope slides and cover-slips, pipette tips and plastic spreaders into the jars of DISINFECTANT immediately after use. NEVER lay pipettes on the bench. 7. DISCARD other used items of equipment to the appropriate container. Contaminated items will then be STERILISED prior to washing. 8. Any ACCIDENT, e.g., spilling of cultures or personal injury, however slight must be reported to the demonstrator at once. When cultures are spilled or tubes etc. dropped and broken, the material must be covered with disinfectant and left for a short time before any attempt is made to clean it up. Following disinfection broken glass must be placed in the receptacle provided. 9. At the END of the laboratory period: a. CLEAR the bench tops. Put all glassware and equipment into the appropriate containers and put cultures for incubation into the correct basket. NOTE: You are responsible for ensuring that your cultures are incubated at the appropriate temperature. You must put them in the correct basket. Any cultures left on the bench at the end of the practical class WILL BE DESTROYED. b. Sponge off the bench top with disinfectant solution. c. WASH your hands with antiseptic soap. d. DO NOT take any cultures out of the laboratory.

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Control of Substances Hazardous to Health The chemicals used in the laboratory are listed below along with the precautions required for their safe handling: Chemicals Inorganic compounds Diluted Chloros disinfectant solution Sodium thiosulfate solution, 1 g L-1 ¼ strength Ringer’s solution diluent Organic compounds Crystal violet GCLP Ethanol Fluorescin Luciferin Methylene blue Teepol Proteins/Enzymes Luciferase Culture Media Baird-Parker agar GCLP GCLP Dichloran rose Bengal chloramphenicol agar M-FC broth MacConkey broth MF-Endo broth Membrane lauryl sulfate medium Lauryl tryptose lactose broth Nutrient broth Nutrient agar Peptone saline diluent Plate count agar Slanetz & Bartley agar (contains sodium azide) Violetred bile dextrose agar Micro-organisms Aspergillus glaucus GMP Bacillus cereus Enterococcus faecalis Escherichia coli Penicillium variabile Rhizopus stolonifera Saccharomyces cerevisiae Serratia plymuthicaNCIMB 4612 Staphylococcus aureus Unidentified organisms from natural habitats GCLP: Good Chemical Laboratory Practice GMP: Good Microbiological Practice

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Precautions to be Observed GCLP GCLP GCLP

GCLP GCLP GCLP GCLP

GCLP

GCLP GCLP GCLP GCLP GCLP GCLP GCLP GCLP GCLP GCLP GCLP GCLP

GMP GMP GMP GMP GMP GMP GMP GMP GMP

Description of Microbiology Laboratory Equipment 1. Laminar Air Flow Cabinet

Use: Prevent contamination of biological samples in the environment by passing the air through a filter. 2. Autoclave

Use: For sterilizing culture media and lab glass wares. 3. Microbiological Incubator

Use: For providing suitable environment for microbiological growth. 5

4. BOD Incubator

Use: To maintain and cultivate the cell and microbiological cultures in certain temperature and humidity. 5. Ultra Low Temperature Freezer

Use: Used to store biological samples such as DNA/RNA, plant samples and insect artifacts, autopsy materials, blood, plasma and tissues, chemicals, drugs and antibiotics. 6. Compound Microscope

Use: To see microorganisms after 50 to 2000 times magnification. 6

7. Fluorescence Optical Microscope

Use: Uses fluorescence and phosphorescence to study properties of organic or inorganic substances. 8. Membrane Filtration Apparatus

Use: Allows the isolation and enumeration of microorganisms passing through membranes specific to microorganisms. 9. PCR Thermocycler

Use: The polymerase chain reaction (PCR) is used for genotyping, cloning, mutation detection, sequencing, microarrays, forensics, and paternity testing. 7

10. Fermenter

Use: It is used for the production of foods and pharmaceuticals through fermentation, and for the identification of microbes. 11. Refrigerated Centrifuge

Use: To separate biological samples through centrifugation under refrigerated temperatures. 12. Hot Air Oven

Use: Uses dry heat to sterilize lab glass wares and small equipment. 8

13. Colony Counter

Use: Used to estimate a liquid culture's density of microorganisms by counting individual colonies on an agar plate, slide, mini gel, or Petri dish. 14. Inoculating Loop

Use: Used in the cultivation of microbes on plates by transferring inoculum for streaking. 15. Bunsen Burner

Use: Used to sterilise pieces of equipment and to produce an updraft that forces airborne contaminants away from the working area. 16. Automatic Pipette

Use: Used in chemistry, biology and medicine to transport a measured volume of liquid, often as a media dispenser. 9

Exp. 2: Preparation and Sterilization of Simple Culture Media Theory The survival and growth of microorganisms depend on available and a favourable growth environment. Culture media are nutrient solutions used in laboratories to grow microorganisms. For the successful cultivation of a given microorganism, it is necessary to understand its nutritional requirements and then supply the essential nutrients in the proper form and proportion in a culture medium. The general composition of a medium is as follows:  H-donors and acceptors (approximately 1-15 g/L)  C-source (approximately 1-20 g/L)  N-source (approximately 0.2-2 g/L)  Other inorganic nutrients e.g. S, P (50 mg/L)  Trace elements (0.1-1 µg/L)  Growth factors (amino acids, purines, pyrimidines, occasionally 50 mg/L, vitamins occasionally 0.11 mg/L)  Solidifying agent (e.g. agar 10-20 g/L)  Solvent (usually distilled water)  Buffer chemicals Microbiological culture media could be classified based on consistency as: 1. Liquid Media or Broth: Cultures in liquid media (or broth) are usually handled in tubes or flasks and incubated under static or shaken conditions. This way, homogenous conditions are generated for growth and metabolism studies,(e.g. with the control of optical density and allowing sampling for the analysis of metabolic products).In such cases no agar is added or used while preparing the medium. 2. Semisolid Media: They are usually used in fermentation and cell mobility studies, and are also suitable for promoting anaerobic growth. Such media are prepared by adding half quantity of agar (1/2 than required for solid medium) i.e. about 0.5% in the medium. 3. Solid Media: It is prepared in test tubes or in Petri dishes, in the latter case; the solid medium is called agar plate. In the case of tubes, medium is solidified in a slanted position, which is called agar slant, or in an upright position, which is called agar deep tube. Solid media are used to determine colony morphology, isolate cultures, enumerate and isolate bacteria (e.g. using dilutions from a mixed bacterial population in combination with spreading), and for the detection of specific biochemical reactions (e.g. metabolic activities connected with diffusing extracellular enzymes that act with insoluble substrates of the agar medium).5-7% agar agar or 10-20% gelatin is added to the liquid broth to make solid media. Sterilization of Culture Media Although sterilization of culture media is best carried out in a steam autoclave at temperatures between 121 to 134°C it has to be recognised that damage is caused to the medium by the heating process. Heat-treatment of complex culture media which contain peptides, sugars, minerals and metals results in nutrient destruction, either by direct thermal degradation or by reaction between the medium components. Toxic products caused by chemooxidation can also be formed during heat-treatment. It is important, therefore, to optimise the heating process so that a medium is sterile after heating but minimal damage is caused to the ingredients of the medium. As a general rule it is accepted that short-duration, high-temperature processes are more lethal to organisms and less chemically damaging than are longer, lower temperature processes e.g. 3 minutes at 134°C is preferable to 20 minutes at 115°C. Sterilization Cycle The sterilization cycle can be divided into its four stages: Stage 1: 20°-121°C Chamber heat-up time

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The chamber heat-up time depends on the efficiency of the autoclave (air discharge/steam input) and the size of the load in the chamber. The time required for this stage is measured with a recording probe located in the air-discharge valve located in the base of the chamber. Stage 2: <100°-121°C Heat penetration time of the medium container The heat penetration time depends mainly on the volume of the individual containers, although the shape and the heat-transfer properties of the containers may affect this stage. The time required for the medium volume to reach 121°C is measured with thermocouples placed in the centre of the innermost container. Volume (ml) in glass bottles Time (min)  100 ml 19 min  500 ml 18 min  1000ml 22 min  2000 ml 27 min  5000 ml 37 min These times assume that agar media have been dissolved before autoclaving. It is also assumed that maximum exposure to steam is possible. Thus although the single l00 ml bottle required 12 minutes to reach 121°C, when placed in a crate with other bottles it required 19 minutes and when placed in the centre of stacked crates it required 30 minutes. Stage 3: 121°-121°C Holding time at the prescribed temperature The holding time at 121°C depends on (i) the number of organisms originally present in the medium (ii) the fractional number of an organism presumed present after heating e.g. N = 0.001 equivalent to one bottle in every 1000 bottles heated becoming contaminated (iii) the thermal death rate constant of the presumed organism present at 121°C. The recommended holding times are:  Temperature (°C): 121 126 134  Time (minutes) : 20 10 3 Stage 4: 121°- 80°C Cool-down time for the chamber to reach 80°C The cool-down time depends on the size of the load in the chamber and the heat loss rate from the autoclave. Water-sprays are used to accelerate cooling in commercial sterilizers but very careful control is required to avoid bottle fracture and the ingress of the cooling spray into the sterilized medium. The latter problem occurs when the vacuum formed in the head-space during cooling sucks contaminated cooling fluid up the thread of the cap and into the bottle. Sterilization checks Culture media autoclaves should be unlagged and of moderate chamber capacity only. Thermal locks on the doors should prevent them opening when the chamber temperature is above 8O°C but even in these circumstances care should be taken to avoid sudden thermal shock when removing glass bottles of hot liquid from the autoclave. When screw-capped containers are placed in an autoclave the caps should be a half-turn free to allow the escape of heated air. When removed from the autoclave the containers should be allowed to cool down in a laminar airflow cabinet. Alternatively screw-capped containers may be sterilized in a jar which is covered by a piece of felt which effectively protects the containers from infection by air-borne microorganisms. Caps are screwed down tightly after the contents have cooled to ambient temperature. All autoclaves should be checked at fixed periods of time to ensure that they are functioning efficiently. Physical measurements should be made on temperature and pressure readings, the quality of the steam 11

should be checked, the efficiency of the 'near-to-steam' air traps in the base of the autoclave should be determined and the safety valves checked. Mandatory inspections of autoclaves as pressure vessels are normally carried out annually by specialists under instructions from insurers of such apparatus. With small laboratory autoclaves this inspection is not mandatory. Chemical indicators will show the temperature reached or exceeded and some will indicate the time held at the specified temperature. Under-autoclaving is usually self-evident because failure to destroy all the bacterial spores naturally present in dehydrated media (the ‘bioburden’) will allow growth to take place in the stored or incubated medium. Failure of sterilization should always be suspected when contamination of prepared media occurs with sporing organisms. Biological indicators of sterilization will demonstrate the ability of the autoclave to destroy bacterial spores. Overheating effects Overheating is a common cause of pH drift, darkening, precipitation, poor gel strength and reduced bacteriological performance. These effects can also be produced if a concentrated 'pool' of ingredients at the bottom of the container is heated. All culture media should be in solution before sterilization. This will reduce the occurrence of Maillard-type reactions (non-enzymatic browning) taking place in the medium. Overheating effects will occur if agar media are allowed to gel in bottles and are later steamed to melt the agar. They will also occur if molten media are held at 50°C for more than 3 hours before use. Agar media with pH values at or below 5.0 are very sensitive to overheating in any form because the agar is hydrolysed and the gel strength fails. It is recommended to sterilize the agar of media of a pH lower than 5.0 separately. Most of the difficulties in culture media sterilization occur when large unit volumes of media (>2 litres) must be processed. The best solution to this problem is the use of a culture medium preparator. These semiautomatic processors, made by New Brunswick and other manufacturers overcome the problem of poor heat penetration of agar by a continuous stirring or agitation of the medium during the heating phase. Such preparators will significantly reduce the time required for sterilization at 121°C or in some models at 134°C. They are strongly recommended because of their high efficiency and minimal damage to culture media. Preparation and Sterilization of Agar Slants Materials and equipment:  distilled water  measuring cylinder  flask  bacteriological chemicals  laboratory scales  chemical spoons  1N NaOH solution  1N HCl solution  pH indicator paper or pH meter  cotton gloves  dispenser  test tubes  test tube caps  test tube basket  slanting stage  autoclave  incubator

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Working Procedure:  Measure the components of the medium (e.g. TSA or nutrients) into a flask containing 9/10 volume of the solvent. Use a clean chemical spoon for every measurement. Dissolve the solid components and fill with the remaining solvent up to final volume. If the medium contains heat sensitive components (like sugars), they must be separately sterilised in solution (e.g. by filter sterilisation), and then mixed with the already sterilised and cooled agar medium.  Close the flask with cotton plug and cover with aluminium foil, put into the autoclave and start a sterilisation cycle. This cycle could be intermitted when the internal temperature has reached 121°C, at that temperature every component (e.g. agar-agar) will be dissolved correctly.  Check the pH of the medium with an indicator paper or with a pH meter and adjust to the proper value with NaOH or HCl solution.  Pour the 60-70°C medium into the dispenser. Add 5-6 mL medium to each test tube, close them with caps and place them into a test tube basket.  Place the tubes into the autoclave and complete a whole sterilisation cycle for 20 min at 121°C.  Put the test tubes onto a slanting stage to let the medium solidify in the test tubes.  Label the slants according to the type of the medium and perform a sterility test: incubate the test tubes at 28°C for 24 hours, and check for sterility.  The prepared media can be stored for 1-2 weeks at 12-15°C, or longer in a refrigerator. (Do not store medium containing agar-agar under 4-5°C as it destroys its structure!) Preparation and Sterilization of Agar Plates Materials and equipment:  distilled water  measuring cylinder  flask  bacteriological chemicals  laboratory scales  chemical spoons  1N NaOH solution  1N HCl solution  pH indicator paper or pH meter  cotton gloves  sterile, empty Petri dishes  Bunsen burner  autoclave  incubator Working Procedure: 1. Prepare a medium as agar slants. 2. Cool the sterilised medium to 55°C. 3. Take out the cotton plug and flame the mouth of the flask over a Bunsen burner, and then pour the medium into sterile, empty Petri dishes (15-20 mL into each Petri dish). 4. Keep the Petri dishes horizontally until the medium completely solidifies. Turn dishes upside-down and stack them up for storage. 5. Label the plates according to the type of the medium and perform a sterility test as before. 6. In case of longer storage, Petri plates must be placed into plastic bags or boxes to avoid drying out. Special considerations during preparation of sterilized media Liquid media which are sterilized in their final containers should be cooled down to room temperature as rapidly as possible. Screw caps should then be tightened. 13

Containers of agar media which have been sterilized should be placed in a 50°C water bath and the medium dispensed as soon as it reaches this temperature, or within a maximum of 3 hours in the bath. The medium should be mixed thoroughly, without bubble formation and aseptically dispensed into sterile containers. Do not expose dishes of agar media to sunlight; it causes excessive condensation on the lids and may cause the formation of inhibitory substances by photo-oxidation. Heat-labile supplements should be added to the medium after it has cooled to 50°C. Allow the sterile supplement to come to room temperature before adding it to the agar medium. Very cold liquids may cause agar to gel or form transparent flakes which can easily be seen e.g. in blood enriched agar. Mix all supplements into the medium gently and thoroughly, then distribute into the final containers as quickly as possible. Blood used for the preparation of blood agar should be as fresh as possible and should have been stored at 28°C (blood must not be frozen). Warm the blood in a 35°C incubator before addition to sterile molten agar base, which has been cooled to 40-45°C. Adequate mixing in a large head-space vessel is essential to ensure aeration of the blood. Poorly oxygenated blood plates are purplish in colour whereas properly aerated blood agar is cherry-red. Defibrinated blood is recommended for use rather than blood containing an anticoagulant. Storage of prepared media The recommended shelf-life of prepared culture media varies considerably. Screw-capped bottles of nutrient broth and agar can be stored for 6 months at low ambient temperatures (12-l6°C). It is important to store all media away from light. Agar plates should be stored at 2-8°C in sealed containers to avoid loss of moisture. DO NOT FREEZE. Fresh media are better than stored media therefore avoid long storage times. Some very labile beta-lactam selective agents have very short active lives and media containing such substances should be used within a few days of preparation. It is good laboratory practice to establish shelf-lives for all prepared media and date-stamp the containers or holders accordingly. Loss of moisture from agar plates is a common cause of poor bacteriological performance. Do not preincubate all plates overnight as a sterility check. Only obviously wet plates require pre-inoculation drying. Ensure that all plates are incubated in a humid environment. Observations: Examine prepared media before inoculation. Look for evidence of contamination, uneven filling or bubbles on surface of agar, colour changes, haemolysis and signs of dehydration such as shrinking, cracking and loss of volume. Discard any defective plates or tubes. Discussion Table of faults and possible causes in media sterilization 1. Wrong pH value Possible Causes: pH test carried out above 25°C. Overheating through prolonged sterilization, remelting or overlong period at 50°C. Incomplete solution of medium. Poor quality water or containers. Dehydrated medium stored incorrectly or beyond the stated shelf-life. 2. Turbidity, Precipitation Possible Causes: Poor quality water or containers. Overheating or prolonged storage at 50°C. pH value incorrect. Incomplete solution. 14

3. Darkening. Possible Causes: Overheating, incomplete solution or pH drift. Presence of phosphate in addition of glucose or other sugars and agar. 4. Soft gel Possible Causes: Agar not in solution, poor mixing, prolonged storage at 50°C. Overheating at low pH values. Error in weighing or overdilution with inoculum or media supplements. pH too low for agar. 5. Poor bacterial growth Possible Causes: Prolonged and excessive heating, incomplete solution. Inhibitory substances in water or containers. Darkening and pH drift.

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Exp. 3: Application of Aseptic Inoculation Techniques in Solid and Liquid Culture Media Objectives To understand the principles of aseptic technique and be able to apply these principles in:  inoculating a slope of agar culture medium  inoculating a liquid culture medium  streaking out bacterial growth on the surface of an agar nutrient medium to obtain separated colonies Theory Aseptic techniques are methods of handling materials that minimise the chances of microbial contamination. They are used when manipulating pure cultures of microorganisms in order to keep them pure and to prevent the environment from being contaminated. These techniques are also used when manipulating sterile products that, because of their instability, cannot undergo a final sterilisation process, e.g. many pharmaceutical products. These techniques are based on the knowledge that all natural materials, including air, water, dust, clothing, skin, soil, faeces etc. contain micro-organisms that may infect cultures, sterile solutions or other sterile items. Aseptic techniques include the following: 1. 2.

3. 4. 5. 6. 7. 8. 9.

10.

11.

Using only sterile equipment when manipulating cultures or sterile materials- e.g. inoculating loops, pipettes, other glassware etc. Use of microbe-impermeable barriers to exclude air-borne microorganisms and to prevent contact with non-sterile materials - e.g. use of cotton wool plugs in tubes and pipettes (note that cotton wool is only an effective microbial filter when it is dry); use of caps on bottles; wrapping of items in paper or aluminium foil or metal boxes. Working close to the Bunsen flame at all times, because the upward draft near the flame reduces the chance of microbes in air from falling into open containers. Passing the neck of open containers, pipettes and other items briefly through the flame at the start and, in some cases, at the end of manipulations. DO NOT FLAME PLASTIC MATERIALS. Opening containers for the minimum time necessary. Holding open containers in a near horizontal position to reduce the open area exposed to vertically falling particles. Cleaning work surfaces with cotton wool or absorbent tissue paper moistened with water or disinfectant solution (don't use a dry cloth as this tends only to disperse dust into the air). Moistening work surfaces with an oil to cause any dust particles that settle to stick to the surface. The careful disposal of unwanted cultures and contaminated equipment to reduce the general level of microbial contamination in the working environment, e.g. place used pipettes immediately in jars containing disinfectant solution. Use of an inoculating/dispensing hood: a. Simple type - consisting of a box with one glass side for viewing the interior and two holes with gloves through which arms can be put to inoculate cultures and manipulate sterile items inside the box with minimum risk of contamination. Usually a germicidal lamp is fitted which is used to sterilise the interior of the box before starting work. In addition, any or all of the aseptic techniques mentioned above should be used where appropriate. b. Lamina air flow work station - provides a work area over which sterile air is blown. It is the most satisfactory environment for dispensing sterile products. Use of a "sterile" inoculating/dispensing room - a room provided with sterile air (filtered) and an ultra-violet lamp in which manipulations can be done with minimum risk of contamination.

Materials (per student):  1 bottle 5 ml nutrient broth  1 nutrient agar slope  1 nutrient agar plate  access to nutrient agar slope of a known bacterial culture 16

Procedure 1. Spread some bacterial growth from the culture provided over the surface of a nutrient agar slope. 2. Transfer some bacterial growth from the culture provided to a tube of nutrient broth. 3. Streak out some bacterial growth over the surface of a nutrient agar plate so as to obtain at least some well separated colonies on the plate after incubation. Various methods of streaking give satisfactory results in experienced hands, but the following method is recommended as being particularly suitable for beginners. a. Label the base of the plate with the name of the organism, the date and your own name. b. Place the plate, base uppermost, on the bench near the bunsen. Loosen the plug/cap of the tube/bottle containing the culture. c. Sterilise the loop, allow it to cool and pick up a small amount of bacterial growth from the culture. d. Pick up the bottom of the Petri dish and spread some bacterial growth over an area 0.5 - 1 cm in diameter (the pool, (1) in the diagram). e. Sterilise the loop and cool it by touching it on the sterile agar. Streak out the growth from the pool as shown in (1) (i.e., 2-3 times from the pool and 2 - 3 times not entering the pool.) Note that by carefully positioning the angle at which the plate is held relative to the incident light it is quite easy to see previous streaks. Between streaking replace the lid on the plate to reduce the likelihood of contamination. f. Sterilise the loop, cool, and streak as (2). g. Repeat as (f) for (3) and (4), (if sufficient space remains). BE CAREFUL not to re-enter the pool with the final streaks. h. Return the plate to its lid and place it, inverted (i.e., agar uppermost) in the incubation basket. 4. Incubate cultures at 25oC for 2 day.

Observation 1. Examine the appearance of your nutrient broth and nutrient agar slope cultures. 2. Examine the colonies on the streak lines with a hand lens.  If all are identical then the culture is probably pure.  If they are not identical the culture is probably impure (i.e. contains more than one kind of bacteria).  Note that where colonies are crowded together they are smaller than those colonies well separated from other colonies. 3. Look for presence of aerial contaminants; i.e. colonies not on the streak lines and usually differing in appearance from those of the culture. 17

Exp. 4: Morphological Examination of Microorganisms Objectives After completing this practical you should be able to:  Set up a microscope to observe microorganisms under bright field illumination  Prepare a 'wet mount' of a microbial culture or food material for microscopic examination.  Recognise and differentiate between bacteria, yeasts and molds when seen through the microscope. Theory Microorganisms may be defined as living organisms that are invisible to the unaided human eye. The smallest distance between two objects that can be resolved by the unaided eye is about 0.1 mm (100 µm). The majority of microorganisms are smaller than this and in order to see them as distinct objects it is necessary to use a microscope. The light microscope (bright field use) can resolve objects down to 0.2 µm in diameter and this includes all bacteria, protozoa, algae and fungi, but not viruses, most of which are smaller and can only be seen with the electron microscope. Bacterial cells are about 1 µm or less in diameter. Cell morphology (e.g. spherical, rod-shaped, helical, filamentous), together with a few physiological tests, will allow the identification of most bacteria to a major group, but identification to genus or species requires extensive testing of physiological, biochemical and/or genetic properties. Fungi are larger than bacteria with cells or hyphae 3 - 5 µm or more in diameter. They include two major morphological types: a. Unicellular fungi or yeasts. b. Filamentous fungi or molds. Identification of genera and species of yeasts involves both morphological and physiological characteristics. However, identification of mold genera and species is almost entirely based on morphology, particularly of reproductive structures. Molds tend to form their fruiting structures on the surfaces of foods from whence growth can be scraped off and examined in a wet mount. To reduce the formation of air bubbles it is advisable to use a fungal mounting fluid instead of water. If it is desired the fungus can be stained by the use of lactophenol cotton blue, but this has the disadvantage of causing some distortion of the hyphae. The structure of the sporophore is an important aid to the identification of molds, but many mold sporophores are too delicate to be easily transferable to microscope slides without their structure being lost. Hence, a number of methods exist for examining mold fruiting structures with the minimum of disturbance. The methods described below refer to agar cultures, but, depending on the nature of the food, can be adapted for examining molds on foods. In particular, the adhesive tape method is widely applicable. Materials (per 4 students) 1. Bottle culture of Bacillus cereus growing aerobically in a nutrient soup (30°, 1 day) 2. Saccharomyces cerevisiae growing in apple juice (30°, 2 day) 3. Penicillium variabile growing on a nutrient jelly (potato dextrose agar medium, 25°, 3 day). 4. Compound microscope 5. Transparent adhesive tape 6. Petri-dish 7. Scalpel blade

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Setting up the microscope 1. Clean the optical surfaces With the lens tissue provided (a lint free material that will not deposit fibres on the glass) gently polish the upper surface of the light source, the upper surface of the condenser, the bottom surface of the objectives and the upper surface of the eyepiece. 2. Adjust the condenser The condenser is correctly adjusted when it focuses an image of the light source in the same plane as the object. Proceed as follows a. b. c. d.

Raise the sub-stage condenser to its highest point. This will bring it close to its correct position. Close the condenser iris. Place an object slide on the stage and focus it with the X10 objective. Hold an inoculating wire so that the loop rests on the surface of the glass disc over the light source and then slowly lower the condenser until the image of the loop is in sharp focus. This procedure yields the optimum position of the condenser with most microscopes. However, with the Olympus CK some further adjustment is required when using the oil immersion objective (see later).

3. Observation with the X10 objective: a. Open the condenser iris to give adequate but not excessive illumination and contrast. b. Generally this corresponds to about 2/3 of the back of the objective illuminated when viewed with the eyepiece removed. 4. Observation with X40 objective: a. With the X10 objective, select the part of the slide to be observed and place it in the centre of the field of view. b. Rotate the nosepiece so that the X40 objective clicks into place. Only slight adjustment of the fine focus control should be necessary to bring the object into sharp focus. c. Adjust the condenser iris as in (iii). 5. Observation with the oil immersion 100x objective: a. With the 10x objective select the part of the slide to be observed and place it in the centre of the field of view. b. Swing aside the lower power objective and place a drop of immersion oil on top of the preparation. c. Carefully rotate the nosepiece until the oil immersion objective clicks into position. Do not swing the 40x objective through the oil. d. Either - if the object can be seen focus it with the fine focus adjustment Or- if the object cannot be seen, use the coarse focus adjustment and slowly raise the stage until the front of the objective almost touches the slide. Watch this adjustment with your eye level with the stage. Then, look down the eyepiece and very slowly lower the stage with the coarse focus adjustment until the object comes into view. Bring the object into sharp focus with the fine focus adjustment. e. Open the condenser iris fully. Contrast can be increased by partially closing the condenser iris, but this decreases the resolution (i.e. ability to see detail) f. Adjust the condenser very slightly until maximum even illumination is obtained. This avoids the danger of breaking the slide or damaging the very expensive oil immersion lens. 19

6. After use remove oil from the oil immersion objective by wiping with lens tissue. If this is not done the oil may dry to leave a film which will cause the image to be blurred. Remove the slide. Turn the nose piece to leave the low power objective in position. Recognition of Bacteria and Fungi Examine B. cereus and S. cerevisiae in unstained 'wet mounts' as described below. a. Put a grease pencil mark on the centre of a slide and place a drop of the culture over it. Lower on a coverslip and immediately (to prevent drying out) examine the slide under the microscope;

b. Set up the microscope in the normal way, except for keeping the condenser iris partly closed in order to obtain the high contrast required to see unstained bacteria; c. Focus the X10 objective on the grease pencil mark and move it to the centre of the field of view; d. Swing in the X40 objective, focus on the grease mark and look at its edge for microbes; e. Observe bacterial cells exhibiting Brownian movement (see below) and look for motile cells (i.e. cells moving in one direction for a distance many times their diameter). Be careful not to confuse liquid currents in the preparation with motility; f. Make drawings to show the shape, arrangements, and dimensions of the cells. Dimensions of specimens under the microscope can be estimated from a knowledge of the total magnification (= objective magnification x eyepiece magnification). Hence, with 40X objective and 10X eyepiece, giving a total magnification of 400X, an image of 1 mm in size represents an object of 2.5 µm. With the 100X oil immersion objective and 10X eyepiece, a 1 mm image represents a 1 µm object. Recognition of Molds Direct examination of Petri dish cultures a. The mycelium of cultures is usually too dense to allow easy observation of fruiting structures. b. However, where the edges of two colonies approach each other a sterile zone is usually left and at the margin of this zone the spore bearing organs can be easily seen. c. With a clean scalpel blade cut out a 1 cm x 1 cm piece of agar either across the junction of the colonies, or from the edge of the colony towards the zone of mature spores.

d. Examine the living fungus under low power of the microscope (or dissecting microscope if available). 20

e. It is not profitable to make drawings at this stage, but it is important to note features that will subsequently be destroyed during preparation of liquid mounts. e.g., whether spore bearing structures stand up from the mycelium or are formed within it; and whether spores are produced singly, in chains, or clustered together in drops of liquid. Agar block method a. Pipette a drop of fungal mounting fluid onto the agar block (from above) and, placing the edge of a coverslip on the youngest side of the colony, lower it down gently like a hinged lid. b. If this is done correctly any air bubbles and detached spores tend to be carried to one side of the preparation. Adhesive tape method a. Put a drop of mounting fluid onto a slide. b. Cut off a piece of transparent adhesive tape about 8 cm long and 1 - 2 cm wide, and lightly touch the surface of the mould growth with the centre part of the sticky side of the tape. Choose a part of the colony on the edge of the zone of mature spores in order to attempt to obtain developing fruiting structures. c. Place the inoculated region of the tape in the drop of mounting fluid and then stick the ends to the slide. d. Examine the preparations under X10 and X40 objectives and make accurate drawings of reproductive structures. Observation 1. Draw and label different parts of a compound microscope.

2. Diagram of B. cereus 3. Diagram of S. cerevisiae 4. Diagram of P. variabile 21

Exp. 5: Gram Staining Method: Identification of Gram Positive and Gram Negative Microorganisms with Physiological Examination Objective To differentiate between two large groups of microorganisms based on their cell wall composition:  Gram positive bacteria: purple stained bacteria after staining  Gram negative bacteria: pink coloured bacteria after staining Theory The Gram stain was devised by the Danish physician, Hans Christian Joachim Gram, while working in Berlin in 1883. He later published this procedure in 1884. It is a staining technique used to classify bacteria. Bacteria are stained with gentian violet and then treated with Gram's solution. After being decolorized with alcohol and treated with safranine and washed in water, those that retain the gentian violet are Gram-positive and those that do not retain it are Gram-negative. The Gram Reaction is dependent on permeability of the bacterial cell wall and cytoplasmic membrane, to the dye-iodine complex. In Gram positive bacteria, the crystal violet dye –iodine complex combines to form a larger molecule which precipitates within the cell. Also the alcohol/acetone mixture which acts as a decolorizing agent causes dehydration of the multi-layered peptidoglycan of the cell wall. This causes decreasing of the space between the molecules causing the cell wall to trap the crystal violet iodine complex within the cell. Hence the Gram positive bacteria do not get decolorized and retain primary dye appearing violet. Also, Gram positive bacteria have more acidic protoplasm and hence bind to the basic dye more firmly. In the case of Gram negative bacteria, the alcohol, being a lipid solvent, dissolves the outer lipopolysaccharide membrane of the cell wall and also damage the cytoplasmic membrane to which the peptidoglycan is attached. As a result, the dye-iodine complex is not retained within the cell and permeates out of it during the process of decolourisation. Hence when a counter stain is added, they take up the colour of the stain and appear pink.

Materials Required  Clean grease-free slide  Bacteria to be stained  Inoculating loops  Bunsen burner  Pencil marker Reagents Required  Primary stain: Crystal violet  Mordant: Gram’s Iodine  Decolourizer: 95% Ethanol or 1:1 acetone with ethanol  Counter stain: Safranin 22

Preparation of Reagents 1. Primary Stain : Crystal violet Solution A: o Crystal violet = 2 gm o Ethyl alcohol= 20 ml Solution B: o Ammonium oxalate = 0.8 gm o Distilled water = 80 ml Mix solution A and B. Keep for 24 hours and filter. Store in an amber coloured bottle. 2. Mordant : Gram’s Iodine o Iodine = 1 gm o Potassium iodide = 2 gm o Distilled water = to 100 ml Mix and Store in an amber coloured bottle. 3. Decolourizer : 95% Ethanol or 1:1 acetone with ethanol o Acetone = 50 ml o Ethanol (95%) = 50ml 4. Counter stain: Safranin o Safranin O = 0.34 gm o Absolute alcohol = 10ml o Distilled water = 90ml Mix, filter and store in amber coloured bottle. Staining Procedure 



Smear Preparation i. Take a grease free dry slide. ii. Sterilize the inoculating loop on a flame of a Bunsen burner. iii. Transfer a loopful of culture (or the specimen) by sterile loop and make a smear at the center. Smear should not be very thin or very thick. iv. Allow the smear to dry in the air. v. Fix the dry smear by any of the following method: a. Heat fixation o Pass air-dried smears through a flame two or three times. Do not overheat. o Allow slide to cool before staining. b. Methanol fixation o Place air-dried smears in a coplin jar with methanol for one minute. Alternatively, flood smear with methanol for 1 minute. o Drain slides and allow drying before staining. Staining i. Place the slides on the staining rods. ii. Cover the smear with crystal violet stain and leave for 1 minute. iii. Wash carefully under running tap water. iv. Flood the smear with Gram’s iodine solution and leave for 1 minute. v. Drain off the iodine Wash the slide for the again in a gentle stream of tap water. 23

vi.

vii. viii. ix. x.

Flood the slide with the decolorizing agent then wait for 20-30 seconds. This can also be done by adding a drop by drop to the slide until the decolorizing agent running from the slides runs clear. Gently wash the slide under running tap water and drain completely. Counter stain with safranin for and wait for about 30 seconds to 1 minute. Wash slide in a gentile and indirect stream of tap water until no color appears in the effluent and then blot dry with absorbent paper. Observe under microscope.

Observation The staining results of gram stain are as follows:  Colour o Gram Positive: Dark purple o Gram Negative: Pale pink to dark red

 

Shape o Spherical – cocci o Rod – bacilli Arrangement o Cocci in clusters – staphylococci o Cocci in chains – streptococci

Identify probable bacteria from the following list: 1. Gram positive cocci in clusters: Staphylococci species. 2. Gram positive bacilli: Clostridium species, Corynebacterium species, Bacillus anthracis. 3. Gram negative cocci in chains: Streptococci species. 4. Gram negative cocci: Neisseria species. 5. Gram negative bacilli: Escherichia coli, Klebsiella pneumonia.

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Exp. 6: Standard Plate Count (SPC) of Food Samples Objective 

To train on bacteria, yeast and mold counting guidelines to calculate CFU/g from different foods.

Theory Viable Plate Count (also called a Standard Plate Count) is one of the most common methods, for enumeration of bacteria. Serial dilutions of bacteria are plated onto an agar plate. Dilution procedure influences overall counting process. The suspension is spread over the surface of growth medium. The plates are incubated so that colonies are formed. Multiplication of a bacterium on solid media results in the formation of a macroscopic colony visible to naked eye. It is assumed that each colony arises from an individual viable cell. Total number of colonies is counted and this number multiplied by the dilution factor to find out concentration of cells in the original sample. Counting plates should have 30-300 colonies at least. Since the enumeration of microorganisms involves the use of extremely small dilutions and extremely large numbers of cells, scientific notation is routinely used in calculations. A major limitation in this method is selectivity. The nature of the growth medium and the incubation conditions determine which bacteria can grow and thus be counted. Viable counting measures only those cells that are capable of growth on the given medium under the set of conditions used for incubation. Sometimes cells are viable but non-culturable. The number of bacteria in a given sample is usually too great to be counted directly. However, if the sample is serially diluted and then plated out on an agar surface in such a manner that single isolated bacteria form visible isolated colonies, the number of colonies can be used as a measure of the number of viable (living) cells in that known dilution. The viable plate count method is an indirect measurement of cell density and reveals information related only to live bacteria. Normally, the bacterial sample is diluted by factors of 10 and plated on agar. After incubation, the number of colonies on a dilution plate showing between 30 and 300 colonies is determined. A plate having 30-300 colonies is chosen because this range is considered statistically significant. If there are less than 30 colonies on the plate, small errors in dilution technique or the presence of a few contaminants will have a drastic effect on the final count. Likewise, if there are more than 300 colonies on the plate, there will be poor isolation and colonies will have grown together. Generally, one wants to determine the number of (colony forming units) CFUs per gram (g) of sample. To find this, the number of colonies (on a plate having 30-300 colonies) is multiplied by the number of times the original g of bacteria was diluted (the dilution factor of the plate counted). For example, if a plate containing a 1/1,000,000 dilution of the original g of sample shows 150 colonies, then 150 represents 1/1,000,000 the number of CFUs present in the original g. Therefore the number of CFUs per g in the original sample is found by multiplying 150 x 1,000,000 as shown in the formula below: 𝐶𝐹𝑈𝑠 𝑝𝑒𝑟 𝑔 𝑜𝑓 𝑠𝑎𝑚𝑝𝑙𝑒 = 𝑛𝑢𝑚𝑏𝑒𝑟 𝑜𝑓 𝑐𝑜𝑙𝑜𝑛𝑖𝑒𝑠 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 × 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟 𝑜𝑓 𝑡ℎ𝑒 𝑝𝑙𝑎𝑡𝑒 𝑐𝑜𝑢𝑛𝑡𝑒𝑑 At the end of the incubation period, select all of the agar plates containing between 30 and 300 colonies. Plates with more than 300 colonies cannot be counted and are designated "too numerous to count" (TNTC). Plates with fewer than 30 colonies are designated "too few to count" (TFTC). Materials Required 1. Work area, level table with ample surface in room that is clean, well-lighted (100 foot-candles at working surface) and well-ventilated, and reasonably free of dust and drafts. The microbial density of air 25

in working area, measured in fallout pour plates taken during plating, should not exceed 15 colonies/plate during 15 min exposure 2. Storage space, free of dust and insects and adequate for protection of equipment and supplies 3. Petri dishes, glass or plastic (at least 15 × 90 mm) 4. Pipets with pipet aids (no mouth pipetting) or pipettors, 1, 5, and 10 ml, graduated in 0.1 ml units 5. Dilution bottles (160 ml), borosilicate-resistant glass, with rubber stoppers or plastic screw caps 6. Pipet and petri dish containers, adequate for protection 7. Circulating water bath, for tempering agar, thermostatically controlled to 45 ± 1°C 8. Incubator, 35 ± 1°C 9. Tally register 10. Dilution blanks, 90 ± 1 ml sterile saline solution 11. Refrigerator, to cool and maintain samples at 0-5°C 12. Freezer, to maintain frozen samples from -15 to -20°C 13. Thermometers (mercury) appropriate range Samples: 1. Bacteria from yoghurt 2. Yeast from bread 3. Mold from bread Working Procedure 1. Take 6 dilution tubes, each containing 9 ml of sterile saline. 2. Dilute 1 ml of a sample by withdrawing 1 ml of the sample and dispensing this 1 ml into the first dilution tube. 3. Using the same procedure, withdraw 1 ml from the first dilution tube and dispense into the second dilution tube. Subsequently withdraw 1 ml from the second dilution tube and dispense into the third dilution tube. 4. Continue doing this from tube to tube until the dilution is completed. 5. Transfer 1 ml from each of the dilution tubes onto the surface of the corresponding agar plates. 6. Incubate the agar plates at 37°C for 48 hours. 7. Choose a plate that appears to have between 30 and 300 colonies. 8. Count the exact number of colonies on that plate. 9. Work similarly for each of the samples to count bacteria, yeast and molds.

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Calculation Calculate the number of CFUs per ml of original sample as follows: CFUs per ml of sample = the number of colonies × the dilution factor of the plate counted Computing and recording counts To avoid creating a fictitious impression of precision and accuracy when computing aerobic plate count (APC), report only the first two significant digits. Round off to two significant figures only at the time of conversion to SPC. For milk samples, when plates for all dilutions have no colonies, report APC as less than 30 colonies estimated count. Round by raising the second digit to the next highest number when the third digit is 6, 7, 8, or 9 and use zeros for each successive digit toward the right from the second digit. Round down when the third digit is 1, 2, 3, or 4. When the third digit is 5, round up when the second digit is odd and round down when the second digit is even.



Plates with 30-300 CFU:

Calculate the APC as follows:

27

Where: o N = Number of colonies per ml or g of product o ∑C = Sum of all colonies on all plates counted o n1 = Number of plates in first dilution counted o n2 = Number of plates in second dilution counted o d = Dilution from which the first counts were obtained

= 537/0.022 = 24,409 ≈ 24,000 When counts of duplicate plates fall within and without the 30-300 colony range, use only those counts that fall within this range 

All plates with fewer than 30 CFU When plates from both dilutions yield fewer than 30 CFU each, record actual plate count but record the count as less than 30 × 1/d when d is the dilution factor for the dilution from which the first counts were obtained.



All plates with more than 250 CFU When plates from both 2 dilutions yield more than 300 CFU each (but fewer than 100/cm2), estimate the aerobic counts from the plates (EAPC) nearest 300 and multiply by the dilution.

o TNTC, too numerous to count. o EAPC, estimated aerobic plate count.

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Exp. 7: Use of Beneficial Microorganism in Producing Fermented Food Product (Dahi) Objective  To produce fermented food product using beneficial microorganisms Theory Fermentation is a metabolic process in which an organism converts a carbohydrate such as starch and sugar into alcohol and/or acid. In 1850s and 1860s Louis Pasteur became the first scientist to study fermentation when he demonstrated that lactic acid fermentation was caused by living organism. Fermented foods are those food produced by modification of raw material of either animal or vegetable origin by the activities of microorganisms. Bacteria, yeast and moulds can be used to produce a diverse range of products that differ in flavour, texture and stability from the original raw material. Different types of fermented products are produced throughout the world:  Cheese  Yogurt  Cultured buttermilk  Acidophilus milk  Curd  Kefir  Shrikhand/Chakka  Lassi Following table shows the different beneficial microorganisms involved in fermented food product manufacture:

Dahi/Curd Curd is well known dairy product obtained by Lactic Acid fermentation of milk. It is generally consumed in its original form as an accompanied to the meal or it may be turned into raita by mixing it with grated cucumber, diced boiled potato, fried bits of gram flour batter, or pulsed based vadas. Curd or Dahi may be consumed as a sweet or savoury lassi drink or as a dessert containing sugar and fresh diced banana, orange 29

slices, mango bits and other seasonal fruits. In India system of medicine (Ayurveda), curd has been strongly recommended for curing ailments like dyspepsia, dysentery and other gastrointestinal disorders. This product is also believed to improve appetite and vitality. Some of the beneficial effects of curd are attributed to the antibacterial components formed during the fermentation and the low pH that prevents the growth of putrefactive and other undesirable organisms including potential pathogens and possesses an increased digestibility. Types of Dahi

Ingredients Required  Milk  Specific starter culture (Lactobacillus lactis)  Sugar Materials required  Pan  Spatula  Stove  Incubator  Plastic cups Manufacturing Procedure

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Observations 

Sensory evaluation Body, taste, colour and flavour



Physico-chemical evaluation Moisture content, total soluble solids (TSS), acidity, pH

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