ACP010: INTRODUCTION TO MICROBIOLOGY FOR DIPLOMA IN … · Microbiology Lab Practices and Safety...

Prof. James W. Muthomi 2016 Department of Plant Science and Crop Protection, University of Nairobi Page 1 of 23

ACP010: INTRODUCTION TO MICROBIOLOGY FOR DIPLOMA IN CROP PROTECTION

LABORATORY PRACTICALS

PRACTICAL 1: TOUR OF THE LAB – PRACTICES, SAFETY RULES, EQUIPMENT AND APPARATUS

1. Objectives

i. To learn the laboratory practices and safety rules

ii. To familiarize with equipment and apparatus used in microbiology laboratory

2. Microbiology Lab Practices and Safety Rules i. Wash hands with disinfectant soap when you arrive at the lab and again before you

leave. ii. Absolutely no food, drinks, chewing gum, or smoking is allowed in the laboratory.

Do not put anything in the mouth such as pencils, pens, labels, or fingers. Do not store food in areas where microorganisms are stored (No foods in laboratory refrigerators; no cooking in the laboratory).

iii. Purchase a long sleeved lab coat and wear it during laboratory sessions. The lab coat must cover the arms and be able to be removed without pulling it over the head. The lab coat should be removed after the practical session and do not wear it to other non-lab areas.

iv. Avoid loose fitting items of clothing. Wear appropriate clothing and shoes (shorts and sandals are not allowed) in the laboratory. Long hair should be ties back to minimize contamination of cultures and fire hazards.

v. Keep the work benches free of all unnecessary materials (Backpacks, purses, coats, sweaters, books, bags, papers,) except the laboratory apparatus and your note book.

vi. Disinfect work areas with 70% ethanol or fresh 10% bleach before and after use. Laboratory equipment and work surfaces should be decontaminated with an appropriate disinfectant on a routine basis, and especially after spills, splashes, or other contamination.

vii. Label everything clearly: Label all plates, tubes, cultures etc before starting an experiment. Indicate your name and date of experiment.

viii. Replace caps on reagents, solution bottles, and bacterial cultures. Do not open Petri dishes unless absolutely necessary.

ix. Inoculating loops and needles should be flame sterilized in a Bunsen burner or spirit lamp before they are laid down on the bench.

x. Turn off Bunsen burners or spirit lamp when not in use. Long hair must be restrained if Bunsen burners or spirit lamp are in use.

xi. When flame sterilizing with alcohol, be sure that there no papers nearby. Never spray ethanol near flame; turn the burner off when not in use.

xii. All microorganisms should be treated as potential pathogens. Use appropriate care and do not take cultures out of the laboratory.

xiii. Never pipette by mouth. Use a pipetting aid or adjustable volume pipettors. Mouth pipetting is strictly prohibited as it has been known to result in many laboratory-acquired infections or poisoning.


xiv. Consider everything a biohazard - do not pour anything down the sink. Autoclave liquids and broth cultures to sterilize them before discarding. Dispose of all solid waste material in a biohazard bag and autoclave it before discarding in the regular trash.

xv. Familiarize yourself with the location of safety equipment in the lab (e.g., eye-wash station, sinks, fire extinguisher, biological safety cabinet, first aid kit, emergency gas valve).

xvi. Dispose of broken glass, razor blades, syringe needles and sharp metal objects in the broken glass or “sharps” container.

xvii. Report spills and accidents immediately to a laboratory technician. Clean small spills with care. Seek help for large spills.

xviii. Report all injuries or accidents immediately to the instructor or laboratory technician, no matter how small they seem.

xix. Materials such as stains, reagent bottles, Petri plates, pipettes, microscopes etc must be returned to their original place after use.

xx. Handled all equipment with care: never open or dismantle parts of equipment. Equipment like ovens, microscopes, etc must be switched off whenever not in use. Microscope stage, lenses and objectives should be cleaned with special lens tissue before and after use.

xxi. Keep the laboratory doors and windows closed when experiments are in progress.

3. Equipment and apparatus used in microbiology laboratory The class will be divided into groups of three to five students. The following will be

demonstrated:

i) The use of the apparatus/ equipment/ glass ware

ii) How to operate the equipment

iii) The handling and care of the equipment, apparatus and glassware

Equipment - Laminar flow hood, microscope, water bath, hot air oven, iIncubator, refrigerator, autoclave, hotplate/Heater, centrifuge, pH meter, spectrophotometer, colony counter, weighing balance, homogenizers or blender, water-bath Apparatus - Tripod, bunsen burner or spirit lamp, transfer needle, inoculating wire loop, dissecting needles, forceps, scissors, ocular micrometer, stage micrometer, burette setup, thermometers, plastic wash bottles, glass rods, L-shaped glass rods, V-shaped glass rods, staining rack Glassware - Petri dishes, conical flasks, culture tubes without screw caps, screw-capped tubes (universal bottles), Durham fermentation tubes, beakers, funnels, graduated cylinders, graduated pipettes, capillary pipettes, dropper bottle for staining reagents, screw-capped bottles for stock reagents, glass microscope slides, depression (concave) slides, glass cover slips


Questions 1. Explain the importance of cleaning hands before leaving the laboratory

2. What is the purpose of wearing a laboratory coat when working in the lab

3. Explain why the wire loops and inoculating needles should be flame sterilized before

placing on the bench after inoculation.

4. Explain the importance of not using ethanol near a spirit lamp or Bunsen burner

5. State the uses of the following equipment:

a. Water bath

b. Blender

c. Hot air oven

d. Auto clave

6. Explain the differences between the following apparatus:

a. Cover slip and microscope slide

b. Wire loop and inoculating needle

c. Test tube and universal bottle

d. Forceps and spatula

e. Conical flask and beaker

PRACTICAL 2: USE AND CARE OF MICROSCOPES 1. Objectives

i. To learn how to identify the parts of a compound microscope. ii. To learn the correct use and care of the compound light microscopes.

iii. To learn how to recording microscopic observations 2. Materials required Magnifying glasses, dissecting microscopes, compound light microscopes, prepared slides of bacteria and fungi, prepared freshly stained bacteria slides, immersion oil, lens paper 3. How to Move and Transport Microscopes

To remove microscopes from the storage area, place one hand completely around the

arm, and then place your other hand underneath the microscope base. Always carry the

microscope in an upright position. Carrying in any other way may allow parts to fall from

the microscope. Use care to ensure that electrical cords are not entangled with those of

other microscopes. Place microscope on a clean area of the desk or laboratory bench.

4. Parts of dissecting and compound microscopes

Two types of light microscopes commonly are used in introductory plant pathology

courses. These microscopes are the compound microscope (Figure 1) and the dissecting

or stereo-microscope (Figure 2). Dissecting microscopes are commonly used for the

observation of larger objects and generally have magnifications of less than 100x


5. Procedures in the use of microscope

i. Turn the microscope on and set the light source on its highest setting. ii. Use the coarse adjustment knob to obtain maximum working distance.

iii. Place the slide on the stage. The slide should fit into the slide holder but is not placed under the slide holder.

iv. Use the stage adjustment knob to move the slide edge of the cover slip bisects the hole in the stage.

v. Move the stage until the specimen is in the middle of the stage. vi. Rotate the scanning objective (4X) into place.

vii. Use the coarse adjustment knob to obtain the minimum working distance. Develop the habit of watching this process to be sure the objective does not crash into the slide.

viii. Look through the oculars. Adjust the light with the iris diaphragm lever on the condenser if necessary. Slowly turn the coarse adjustment knob until the edge of the cover slip comes into focus. Use the fine adjustment knob to sharpen the focus.

ix. Use the stage adjustment knob to locate the specimen on the slide. x. Rotate a higher power objective (10X) into place. Use the fine adjustment knob to

sharpen the focus. Do not use the coarse adjustment knob. Adjust the light using the iris diaphragm lever if necessary. The image is now magnified 100X (10X ocular x 10X objective = 100X magnification). Draw what you see under the microscope. Repeat this procedure with objective X40.

xi. When using the high power objective (100X) use the following procedure. a. Rotate the turret halfway between the 40X and 100X objective. b. Place a drop of immersion oil on the slide and rotate the oil immersion

objective (100X) into place. The objective should be immersed in the oil on the slide.

c. Use the fine adjustment knob to sharpen the focus. Adjust the light using the iris diaphragm lever if necessary. Never use the coarse adjustment knob with high power.

Xii When you are finished with the microscope rotate the nose piece to objective x4 and then clean the microscope, as described below, and return it to storage.

http://www.apsnet.org/edcenter/K-12/NewsViews/Pages/DissectingMicrosocpe.aspx

http://www.apsnet.org/edcenter/K-12/NewsViews/Pages/CompoundMicroscope.aspx


6. Procedure for cleaning a microscope 1. Turn off the light and unplug the cord. Store the cord appropriately. 2. Using the coarse adjustment knob to obtain maximum working distance and remove

the slide from the stage. 3. Using lens paper clean all the lenses starting with the cleanest first—oculars, 4X

through 100X objectives. 4. Clean any oil off of the stage using paper towels. 5. Rotate the scanning objective into place. Use the coarse adjustment knob to obtain

minimum working distance. 6. Return the microscope to the appropriate storage area

Questions

1. List the difference between the dissecting and compound microscope

Dissecting Compound

1

2

3

4

5

2. Name the parts of microscope on the diagram below:

3. State the purpose of each of the following components of a compound microscope:

a. Fine adjustment knob

b. Course adjustment knob

c. Iris diaphragm


d. Mechanical stage control

e. Condenser

4. What is the purpose of adding immersion oil when using the 100X objective?

5. Fill in the table below (5 marks)

Magnification

Objective Objective Ocular Total

Scanning

Low power

Medium power

High power

Oil immersion

6. Make drawing of the objects you have observed under the microscope and indicate the magnification (5 marks)

7. Explain the purpose of oil when using objective X100 ( 3 marks)

PRACTICAL 3: ASEPTIC PROCEDURES IN MICROBIOLOGY 1. Objectives

i. To familiarize with the aseptic followed in the study of microorganisms ii. To learn how to sterilize working surfaces, glassware and solution used in the study

of microorganisms

2. Aseptic techniques i. Disinfect the tables with 70% ethanol before use. ii. Wear gloves and lab coats. iii. When labeling Petri plates, always write on the bottom of the plate. iv. When using hot air oven, give them ample time to warm up before sterilizing loops or

needles. DO NOT leave loops or needles unattended in the oven. They will melt! You will burn your fingers!

v. When inoculating cultures, always sterilize the loop or needle before going into a culture and after transferring it. Sterilize the loop even if you are going back into the same culture again.

vi. Make sure you let your loop cool first – you don’t want to kill the microorganisms. vii. After removing the lid of a test tube, flask or universal bottle, briefly flame the mouth of

the tube/flask before pouring liquids or inserting an inoculating loop, and flame again before replacing the cap.


viii. Don’t place sterile apparatus (loops, needles, pipettes, test tube lids, etc) on the table. Once they touch the table they are no longer sterile.

ix. Sterilized pipettes should remain in canister until just before use. x. Do not open or expose culture media to the air unnecessarily – bacteria and fungi in the

air can contaminate the media. xi. Don’t over-inoculate! It doesn’t take much inoculum to start a culture. Simply touch the

loop or needle to the bacterial or fungal growth and obtain a small amount on the loop or needle. Do not “scrape” the culture, and don’t dig into the agar.

xii. When streaking onto an agar plate or slant, make sure the loop doesn’t break the surface of the agar. A gentle gliding motion is all that is necessary to distribute the bacteria on the plate.

xiii. When finished, disinfect the tables again with antibacterial cleaner. Dispose of gloves in the biohazard trash, and remove the lab coat (for laboratory workers, the lab coat is placed in a designated place). Wash your hands!

3. Sterilization of working surfaces and apparatus

Requirements

1. Bunsen burner or spirit lamp, 70% ethanol in hand sprayer, absolute ethanol, paper towels,

plates containing water agar media

2. Transfer needles, inoculating wire loops, dissecting needles, forceps, scissors, glass rods, L-

shaped glass rods, V-shaped glass rods,

Procedure

NOTE: Ethanol is highly flammable and caution should be taken when handling it – no spraying

70% ethanol near flames, do not swab hands with 70% ethanol when near naked flames; restrain

long hair and no flowing clothing.

i. Make sure the working surface if free of books, paper, bags – only the apparatus to use

ii. Apply 70% ethanol on absorbent paper towel and swab hands and the working surface. Allow the

ethanol to evaporate.

iii. Put the Bunsen burner or spirit lamp and ensure it is not next to ethanol

iv. Dip wire loop/ needle/ glass rod/ forceps in absolute ethanol (ensure the handles or parts to be held

are free of ethanol to avoid flames following the ethanol and burn your hand).

v. Remove the item in (4) above from the ethanol and flame off ethanol on spirit lamp or Bunsen burne.

Heat the wire loop/ inoculating needle in a slanting position , ensuring the tip touches the blue part of

the flame. Remove when red-hot (Do not return the hot implement into the ethanol!).

vi. With the left hand, slighthly open a plate containing water agar media and cool the red-hot wire loop or

needle by touching the surface of the agar medium. The needle/ wire loop is now sterile and ready for

use.

4. Sterilization of glassware and solutions

Requirements

i. Hot air oven and autoclave

ii. Aluminium foul, Peteri dish canisters, pipette canisters, glass peteri dishes, glass pipettes and other

appropriate glassware


Procedure

This will be a demonstration and class will be divided into groups of 3-5 students. The following will

be demonstrated:

a) Operation of hot air oven – for sterilization of glass ware and metal apparatus i. Glass ware to be sterilized must be clean and dry

ii. Place glassware (Peteri dishes and pipettes) in metal aluminium canister; small pieces of apparatus such as glass microscope slides, foreceps, spatula etc are wrapped in aluminium foil.

iii. Then, the materials are arranged in oven to ensure uninterrupted air flow. iv. Allow the oven and the materials to attain the required temperature, e.g.160 oC and

allow to heat for 2 hours and switch off the oven. v. Before opening the oven, the temperature is allowed to fall to about 40 oC

b). Operation of autoclave – for sterilization of liquids and culture media

i. The containers should not be completely filled (leave about 2 -3 cm) with the solution to be

sterilized.

ii. The screw caps should be loose to allow escape of air and steam during sterilization and

entry of air during cooling.

iii. Small pieces of glass ware (glass rods, spatula, microscope glass slides, scalpels) should be

wrapped in aluminium foil.

iv. Ensure the autoclave has adequate water covering the heating elements and to generate

steam.

v. Place the materials in the autoclave, close the door and lock the screws.

vi. Switch on as guided, set temperature to 121 oC and timing to 15 minutes; for manual

operated autoclaves, allow temperature to increase to 121oC before stating to time the

duration of sterization.

vii. After sterilization, allow to cool to 45 oC before opening the door – this allows reduction of

pressure and cooling of the contents to a save temperature and pressure.

PRACTICAL 4: PREPARATION OF GROWTH MEDIA FOR ISOLATION OF MICROORGANISMS

1. Objectives

i. To learn the techniques and procedures of preparing microbial culture media from locally available

ingredients

ii. To learn the steps in preparing general purpose culture media using commercially available brands

2. Required

i. Ingredients for making culture media: commercial media (nutrient agar and potato dextrose agar), agar, distilled water, fresh potato tubers, glucose (or ordinary sugar), fat free beef, mushroom.

ii. Apparatus and reagents – media bottles, weighing balance, hot plate, magnetic stirrer, beakers, graduated cylinders, pipettes, universal bottles, sterile Petri


dishes (glass or plastic), autoclave, water bath, 70% ethanol, spirit lamps, absorbent paper towel rolls, cheese cloth.

3. Preparation of potato dextrose agar (PDA) from local available ingredients Ingredients for potato dextrose agar for growth of fungi

Potato 200 g

Dextrose (may use ordinary

sugar)

20 g

Agar 20 g

Water 1,000 ml

Procedure

i. Peel potato and cut into small cubes

ii. Weigh 200 g of the potato cubes

iii. Boil the potato in 1,000 ml of water until soft (easily penetrated by a glass rod)

iv. Allow to cool and strain through cheese cloth into a clean beaker

v. Add 20 g of dextrose (or ordinary sugar) and 20 g agar to the potato extract and heat

the mixture on a hot plate until agar dissolves

vi. Place the mixture in a media bottle and make up volume to 1000 ml

vii. Autoclave at 121 0C for 15 minutes

viii. Allow the media to cool to 45-50 oC (flask can be comfortably held by hand) in a water bath

set at 45 oC and pour 20 ml of the media into each of the Petri dishes (the procedure of

pouring the media to Petri dishes will first be demonstrated)

ix. Allow the media to cool and harden for at least 12 hours before use – the agar plates are ready for the

growing microorganisms.

4. Preparation of nutrient agar (NA) from local available ingredients

Ingredients nutrient agar for growth of bacteria

Fat free beef 300 g

Potato 200 g

Mushroom 100 g

Sugar 2.5 g

Agar 20 g

Water 1,000 ml

Procedure

i. Cut the beef into small pieces and medium sized, peeled potato tuber in four pieces;

cut the mushroom in half

ii. Weigh 300 g of beef, 200 g of potato and 100 g of the mushroom

iii. Add the cut pieces into a beaker containing 1000 ml water and cook for 2 hours

iv. Allow to cool and filter the extract through cheese cloth;


v. Place the extract on a hot plate and while stirring, add agar and sugar until all the

agar is dissolved

vi. Place the solution in a media bottle; make the solution to 1000 ml and autoclave at

121 0C for 15 minutes; the screw cap should be loose before putting in autoclave

vii. Allow to cool to 45-50 oC (flask can be comfortably held by hand) in a water bath set at 45 oC and pour 20 ml of the media into each of the Petri dishes

viii. Allow the media to cool and harden for at least 12 hours before use – the agar plates are ready for the

growing microorganisms

5. Preparation of agar plates from commercially available potato dextrose agar (PDA) and nutrient agar

(NA) culture media

i. Weigh the appropriate amount of each of the commercial PDA and Nutrient agar as per the

manufacturer’s recommendation

ii. Dissolve each separately in 1000 ml distilled water in water bottles on a hot plate; ensure the mixture is

stirred with magnetic stirrer until the agar is dissolved

ix. Autoclave at 121 0C for 15 minutes

x. Allow to cool to 45-50 oC (flask can be comfortably held by hand) in a water bath set at 45 oC and pour 20 ml of the media into each of the Petri dishes

xi. Allow the media to cool and harden for at least 12 hours before use – the agar plates are ready for the

growing microorganisms

6. Preparation of agar slants of potato dextrose agar (PDA) and nutrient agar (NA) (used for storage of

microorganisms)

In this section, use part of the media solutions prepared above but before the autoclaving stage

i. The media ingredients should be prepared and mixed as described in 1, 2, and 3 above

ii. Dispense 10 ml of each media suspension in each of the screw-capped tubes (universal bottles)

iii. Loosely cap the tubes

iv. Autoclave at 121 0C for 15 minutes

v. Allow to cool to about 50 oC (comfortable hand hot)

vi. Place the tubes in a slanting position and allow to solidify and tightly close the screw caps

PRACTICAL 5: ISOLATION AND CULTURE OF MICROORGANISMS

1. Objectives

i. To isolate microorganisms from different environments

ii. To isolate microorganisms from soil by using serial dilution and pour plate method

2. Requirements Potato dextrose agar plates, autoclaved molten PDA media, universal bottles each containing 9 ml sterile distilled water, 70% ethanol, sterile pipettes, sterile Petri dishes, absorbent paper towels, spirit lamp or Bunsen burner, labels or marker pens

3. Isolation of microorganisms from various environments

Each group of 3-5 students will be provided with 15 plates of potato dextrose agar media


Procedure

i. Expose two petri dish of PDA for about five minutes to the air in the laboratory. ii. Expose two petri dish of PDA for about five minutes to the air outside the laboratory iii. Using a spatula, collect soil from outside; decontaminate hands with 70% ethanol; let ethanol evaporate;

pulverize a pinch of the soil between your index finger and thumb and sprinkle a little bit on the surface of two plate containing PDA.

iv. Collect some dead and rotting leaves or other vegetation from outside; decontaminate hands with 70% ethanol; let ethanol evaporate; cut the leaves into small fragments, and place five pieces on two plates containing containing PDA.

v. Collect some actively growing young leaves from plants or grass outside the laboratory; decontaminate hands with 70% ethanol; let ethanol evaporate; cut the leaves into small fragments, and place five pieces on two plates containing PDA.

vi. Without prior decontamination of your hands, open a plate of PDA and touch the surface of the agar media with your fingers; then replace the lid; repeat this with a second plate.

vii. Control: Leave two plate of PDA undisturbed – no opening or touching with hands without first decontaminating with 70% ethanol. Incubate it at room temperature with the other plates.

Label the petri-dishes to indicate the kind of treatment, group number and the date. Incubate the plates at room temperature and examine regularly during the week.

2. Isolation of microorganisms from soil by serial dilution and pour plate method

Procedure: i. Label six universal bottles containing 9 ml of sterile distilled water each as 10-1, 10-2, 10-3, 10-4, 10-5 and

10-6 . ii. Weigh 1 g of pulverized soil and dissolve in 9 ml of sterile distilled water in a universal bottle; mix by

shaking vigorously for about five minutes. iii. Take 1ml of soil suspension and add into the universal bottle labeled 10-1; mix the contents by shaking. iv. From the 10-1, transfer 1ml of the suspension to the bottle labelled 10-2 with a sterile and fresh pipette.

This dilutes the original suspension by 100 times (102). v. From the 10-2 suspension, transfer 1ml of the suspension to the bottle labelled10-3 using a fresh sterile

pipette, thus diluting the original suspension by 10-3 vi. Similarly repeat this procedure for the bottles labeled 10-4, 10-5 and 10-6 till the original sample is diluted

1,000,000 (10-6) and for each step use a fresh sterile pipette. vii. From each of the dilutions 10-1, 10-2, 10-3, 10-4, 10-5 and 10-6, take 1ml of the suspension and place each

of the 1 ml suspension onto sterile Petri dishes correspondingly labeled 10-1, 10-2, 10-3, 10-4, 10-5 and 10-

6. Two Petri dishes should be used for each dilution. viii. Into each Petri dish, gently add about 15ml of the molten PDA agar medium cooled to 450C and gently

swirl to mix the soil suspension throughout the medium (Do not shake vigorously as this results in formation of air bubbles on the media surface).

ix. Allow the plates to solidify and incubate in an inverted position for 3 -7 days at room temperature


NB: Preserve all the cultures obtained during this practical: Microorganisms isolated from

practical 5 will be used during practical 6

Questions 1. With the help of the colony morphology illustration given below, describe the shape, size,

edge and colour of the colonies developing from the isolation plates.

Present your results using a table as below:

Microorganism type shape size edge colour

A

B

C

D


E

2. Based on the results of the isolation plates, where may microorganisms be found?

3. Should plates containing media or cultures be opened and exposed to air? Explain.

4. Based on the results you obtained from isolation plates, should you touch the agar plates

with your hands? Explain.

PRACTICAL 6: PREPARATION OF PURE CULTURES OF MICROORGANISMS AND EXTRACTION OF NEMATODES FROM SOIL

Cultures obtained from practical 5 will be used during today’s practical

1. Objectives

i. To prepare pure cultures of fungi from a mixed culture

ii. To prepare pure cultures of bacteria from a mixed culture

iii. To extract nematodes from soil

2. Requirements

Mixed cultures (cultures containing many different types of microorganisms), freshly collected soil

from plant root zone, plates of PDA medium, plates of NA medium, wire loops, inoculating needles,

facial tissues (or soft tissue paper towels), spirit lamp or Bunsen burner, wash glasses or empty

transparent plastic Petri dishes, droppers, sieves, shallow pans (or plastic bowls)

3. Preparation of pure culture of bacteria by streak-plate method

1. Label all the plates on the bottom.

2. Sterilize a wire loop on spirit lamp flame and allow it to cool

3. Holding the wire loop in your right hand, pick bacterial growth from a well-isolated colony in a mixed

culture from the previous practical.

4. Lift the Petri dish cover with the left hand and open by holding the cover at an angle of about 600 to the

bottom plate

5. Place the loop containing the bacteria inoculum on the agar surface at the edge on the left hand side of

the plate and gently streak from side to side in parallel lines across the surface of the area.

6. Flame the wire loop again and cool by touching the agar at the middle of the agar plate; turn the Petri dish

at 900 . and make another group of parallel streaks perpendicular the first; again, flame the loop, allow to

cool and make a third group of streak perpendicular to the second group.

7. Replace the lid of the Petri dish and incubate the plate in an inverted position at 25 0C for 48-72 hours.

8. Examine the plates for growth of bacterial colonies.

9. Choose a well-isolated colony from where the third group of streaks had been made; using a sterilized

wire loop, touch the bacterial growth.

10. Lift the lid of the agar plate at 450 and inoculate by making parallel streaks on the agar surface.

11. Incubate for 24 hours and observe growth of bacteria


NB: (i) Avoid pressing the loop too firmly against the agar surface as this will damage the agar surface; (ii)

inoculating loop should be cooled by touching the agar surface at the middle of the plate away from the set of

streaks; (iii) the Petri dish lid should never be lifted completely; (iv) the agar media should be prepared 24 hours

before plating to ensure complete drying of the agar surface.

4. Preparation of pure culture of fungi

i. Flame an inoculating needle to red hot on a spirit lamp flame and cool it by touching the surface of a

fresh agar plate.

ii. Using the sterile inoculating needle, cut a very small agar block containing fungal growth at the growing

edge of a well separated colony.

iii. Lift the lid of a fresh agar plate at 450 and place the agar block at the middle of the fresh agar medium

surface.

iv. Incubate the cultures at 250C for 48-72 hours (for bacteria) or for 7-14 days (for fungi).

5. Extraction of nematodes from soil i. Collect fresh, moist soil from around plant roots

ii. Wrap a small handful amount of soil in two layers of facial tissue

iii. Place the wrapped soil on a sieve (mesh or screen) in a small dish. Add water so that

the mesh is slightly covered with water and the soil contacts the water.

iv. Let sit 1-3 days to allow the nematodes to crawl out of the soil. Make sure the

sample stays in contact with the water - do not let the dish become dry. Covering the

dish with plastic wrap or foil will help prevent drying.


v. Remove the bundled soil from the dish

vi. Using a droper, transfer a few drops of the water and place on a Petri dish and

observe the water in the dish using a binocular dissecting microscope to observe the

nematodes.

NB: Preserve all the cultures obtained during this practical: Microorganisms purified

during practical 6 will be used in practical 7 and 8.

PRACTICAL 7: PREPARATION OF BACTERIAL SMEAR, METHODS OF STAINING AND OBSERVATION OF BACTERIA UNDER MICROSCOPE

1. Objectives

i. To learn the procedures of preparing bacterial smear

ii. To learn the methods and procedures of staining bacteria

iii. To study the morphology of bacteria by observation under compound microscope

2. Requirements

24-hr old cultures of Bacillus Spp., Xanthomonas campestris p.v campestris and mycobacteria; Staining

solutions (methylene blue, Nigrosin, crystal violet, Gram’s iodine solution, 95% ethyl alcohol, safranin, carbol

fuchsin); Clean glass slides, Wire loops, Spirit lamps or Bunsen burner; Blotting paper, Staining racks,

Microscopes with X100 objective; Immersion oil, Wash bottles with sterile distilled water; droppers.

3. Preparation of a bacterial smear

A smear is a thin film of microorganism spread out on a microscope slide. The smear is air dried and then

passed with smear side up, through a flame 2 or 3 times to heat fix the bacteria. Heat fixing denatures bacterial

enzymes, preventing autolysis and also enhances the adherence of bacteria to the slide. The preparation is then

ready for staining procedures.


Procedure

i. Take a clean glass slide and wipe with 70% ethanol and let ethanol evaporate. ii. Flame a wire loop to red hot in spirit lamp flame and allow to cool. iii. Put a loopful of sterile distilled water on the cleaned glass slide. iv. Using the wire loop, transfer a small amount of bacterial growth from one of the colonies grown on

nutrient agar into the sterile distilled water droplet. v. With the loop, emulsify the bacterial growth with the water droplet and thinly spread (or smear) over the

glass slide surface. vi. Allow the smear to air dry at room temperature – moving the glass slide in the air will enhance the

drying. vii. Fix the smear by passing rapidly through the tip of the blue portion of the spirit lamp flame 4 to 5 times

(do not burn the smear – the glass should remain comfortable to hold with fingers). viii. Allow the slide to cool; the smear is ready for staining.

The smear appears as a thin, semi-transparent, whitish layer or film fixed to the glass surface.

4. Negative stainining In negative staining the bacterial cells do not interact with the dye and therefore, only the background is stained

while the bacterial cells remain colourless.

Procedure

i. Place one drop of nigrosin at one end of a clean glass slide. ii. With the help of a sterile wire loop, transfer a loopful of bacterial growth and mix with the drop of

nigrosin. Then add a loopful drop of water and mix. iii. Take another clean slide, place it against the drop at an angle of 30 0C and allow the droplet to spread

across the edge of the top slide. iv. Spread the mixture of the stained bacterial suspension out into a thin wide smear by pushing the top

slide to the left along the entire surface of the bottom slide. v. Allow the smear to air dry. vi. Examine the preparation under oil immersion objective. Note the shape, arrangement and size of cells.

Note: Do not spread the drop during mixing it with bacteria; the thickness of the smear should be

uniform; never heat fix the smear.

5. Methylene blue staining i. Take clean glass slides, swab with 95% ethanol using absorbent paper and air dry ii. Following the procedure described in (3) above, make a smear of the available bacteria on a slide. Fix

the smear by first allowing it to air dry and then passing it gently over a flame. iii. Place on staining rack and apply a few drops of Methlene blue stain on the smear and let the stain act

for 1 min. iv. Pour off the stain and wash the smear gently with slow running tap water water. v. Blot dry with absorbent paper (Do not wipe the slide). vi. Examine under oil immersion.

6. Gram Staining

i. Clean a glass slide by swabbing with ethanol using an absorbent tissue. ii. Make a thin smear of the available bacteria as described in (3) above. iii. Let the smear air dry iv. Heat fix the smear by passing over a flame. v. Place the slide on a staining rack. Stain by adding few drops of crystal violet and allow to stand for 1

min; Using a wash bottle, wash off the stain with distilled water for a few seconds.. vi. Flood the smear with Gram’s iodine for 1 min then wash off the iodine solution under tap water.


vii. Add ethyl alcohol drop by drop, until no more colour flows from the smear. viii. Wash the smear with distilled water and drain. ix. Apply safranin to smear for 30 seconds (counter staining). x. Wash with distilled water and blot dry with absorbent paper. Let the stained slides air dry and examine

under oil immersion with x100 objective. xi. Identify the Gram reaction of each bacterium; make sketches for morphology of the bacterium; describe

the morphology and arrangement of cells.

Gram positive bacteria takes the colour of crystal violet (stained violet to purple) and the Gram negative bacteria takes up the colour of the safranian counter stain (red).

7. Acid-fast staining

i. Prepare a smear of the bacterium on a slide. ii. Air dry and heat fix over a flame. iii. Flood the smear with carbol fuchsin and gently heat (not boil) over a flame for 3 to 5 minutes. From

time to time, add more stain to prevent the smears from becoming dry. iv. Cool the slide and then wash off excess stain with distilled water. v. Decolourize the smear with acid-alcohol until all red colour is removed. vi. Wash with distilled water. vii. Counter stain with methlene blue for 1-2min. viii. Wash with distilled water and blot dry with absorbent paper. ix. Observe under oil immersion at objective x100.

Record the colour of the test bacterium (acid-fast or non-acid-fast); describe the morphology and

arrangement of cells. Non acid fast bacteria stain a deep blue. Acid fast bacteria remain red, being

saturated with the red carbolfuchsin).

PRACTICAL 8: PREPARATION OF MOIST CHAMBER AND MICROSCOPIC STUDY OF FUNGI

Objectives

i. To learn the techniques of inducing formation of fungal structures by the use of moist chamber

ii. To learn the procedures of using dissecting microscope in the study of fungi

iii. To learn the procedures of making microscopes slide mounts of fungi

Requirements

14-21 day-old cultures of fungi, Clean glass slides, Cover slips, Mounting needles, Distilled water, Cotton blue in

lactophenol in a dropper bottle, Spirit lamp or Bunsen burner, 70% alcohol, Microscopes.

Preparation of moist chamber for growth of fungi

The simplest method of growing fungi or molds is to put a substrate in a moist chamber. The substrate provides nutrients, and the chamber maintains the high humidity that favours the growth of fungi. Materials used in preparation of moist chambers include plastic sandwich bags or plastic sandwich box, or Petri dishes.


Requirements: - diseased plant materials (dry vegetation, bread, fruits, leaves with leaf spots, coffee berries with coffee berry disease), "zip lock" sandwich bags with plant or sandwich boxes, paper towel, distilled water, marker pen or labels.

Procedure

i. Label the bags or sandwich box with your name and type of material to be incubated.

ii. Place a damp paper towel in each bag. iii. Place the diseased plant material or other substrate on top of the damp towel. iv. Seal the bags or the box. v. Place the bags in a warm area out of direct sunlight where they will not be disturbed.

vi. Check the bags each day. Fungal growth should be visable in 3 to 5 days. Fungi are fuzzy or hairy and may be green, white, black, yellow, etc.

vii. Record the number, color, and size of the fungal colonies. In the case of bread and some other materials, Rhizopus spp may completely cover substrate in just a couple of days. The fungal growth can now be observed under dissecting microscope or small fragments transferred on microscopic slide, stained and observed under compound microscope.

Use of dissecting microscope in study of fungi

A dissecting microscope is used to examine diseased material for the presence of small fungal structures, such as spores and hyphae under low magnification (up to approximately ×100). Using the dissecting microscope, such structures can be easily transferred to a slide preparation for examination under a compound microscope, at higher magnification (up to ×400). Requirements: Pure cultures of fungi, materials incubated in moist chamber, dissecting microscope, inoculating needles, plates of PDA medium Procedure

i. Using the focussing adjustment knobs, lower the objective of dissecting microscope to the lowest point

ii. Place an open culture plate or plant materials (from moist chamber) containing fungi and place under the objective

iii. Focus by slowly moving the objective upward until the fungal growth is clear iv. Locate the spores or spore-forming structures and hyphae of the fungus – make

diagrams of your observations. v. While observing the material under the microscope, collect spores or small pieces of

mycelia from the fungal growth using a sterile inoculating needle. vi. Transfer the spores or mycelia onto centre of a plate of PDA medium; place a drop of

lactic acid at the edge of the plate (lactic acid reduces the pH and suppresses bacterial growth); incubate the plates to allow growth of the fungus.

Temporary wet mounts for microscopic observation of fungi


Requirements: Pure cultures of fungi, materials incubated in moist chamber, compound microscope, inoculating needles, microscope glass slides, cover slips, distilled water, droppers, cotton in lactophenol stain

Procedure

i. Use materials prepared in moist chamber and pure culture of fungi you prepared earlier ii. Place a small drop of distilled water on the centre of a clean glass slide

iii. Using sterile inoculating needle, transfer a small amount of fungal growth onto the drop of water on glass slide

iv. Place a coverslip with one side touching the slide near one edge of the water drop. v. Gently lower the other side of the coverslip onto the water drop—this method excludes

air bubbles from the preparation (A gentle pressure can be applied to the cover slip by tapping with back of mounting needle or pencil in order to spread the fungal structures and expel air bubbles).

vi. Use a strip of blotting or filter paper to blot excess water at the edge of the coverslip. vii. Examine the preparation under low –power (X10 and X20) and high-power (x40)

objectives.

The procedure may be repeated using cotton blue in lactophenol stain

Note the size, shape and characteristics of the fungus (spores, hyphae);describe the type of

hyphae, spores and conidiophores. Draw sketches of the observed fungal structures.

PRACTICAL 9: MICROSCOPIC STUDY OF MORPHOLOGY OF DIFFERENT GROUPS OF FUNGI

1. Objectives

i. To observe the types of hyphae and spores formed by different fungi

ii. To observe complex mycelium and spores formed by the mushroom

2. Requirements

Cultures of fungi, materials incubated in moist chamber, dissecting microscope, compound microscopes,

microscope slides; cover slips, inoculating needles, scalpel blades, distilled water, cotton blue in lactophenol,

spirit lamp or Bunsen burner,

3. Materials for examination

i. Materials incubated in moist chamber: potato or tomato leaves infected with late

blight, bread, maize leaves infected with northern leaf blight, avocado or ripe banana

with anthracnose rot

ii. Fresh disease plant materials – powdery mildew, coffee rust.

iii. Cultures – Pythium, Fusarium, Colletotrichum, Sclerotinia or Macrophomina with

sclerotia,


iv. Mushrooms

4. Examination under dissecting microscope i. Place an open culture plate or plant materials (from moist chamber) containing fungi

and place under the objective ii. Focus and locate the spores or spore-forming structures and hyphae of the fungus –

make diagrams of your observations.

5. Examination under compound microscope

i. Use materials prepared in moist chamber and pure culture of fungi you prepared earlier

ii. Place a small drop of distilled water on the centre of a clean glass slide iii. Using sterile inoculating needle, transfer a small amount of fungal growth

onto the drop of water on glass slide iv. Place a coverslip and blot excess water at the edge of the coverslip. v. Examine the preparation under low –power (X10 and X20) and high-power

(x40) objectives.

Repeat the procedure using cotton blue in lactophenol stain

Note the size, shape, septation and other characteristics of the fungus (spores, hyphae);

describe the type of hyphae, spores and conidiophores. Draw sketches of the observed

fungal structures.

6. Examination of complex tissues of mushroom under microscope

i. Cut out the stem of the mushroom and pinch it between fingers until it breaks into two or

more long pieces

ii. Gently pull the pieces apart and place the stem sections under the dissecting microscope

and examine the hyphae – draw and label the parts you see.

iii. Using a scalpel, cut through the mushroom cap;

iv. Using a forceps, carefully remove one gill from the cap (avoid touching the free end of the

gill) – grasp the gill near where it attaches to the cap

v. Place the gill on a microscope slide in a drop of water; apply cover slip

vi. Observe under low power (5X, 10X and 20X objectives) – observe finger-like projections on

the surface of the gill (these are spore-bearing structures called basidia). Can you see the

spores? Make drawings of your observation


PRACTICAL 10: DETERMINATION OF POPULATION OF MICROORGANISMS

1. Objectives i. To learn how to use of a haemocytometer ii. To learn the procedure of calculating the concentration of fungal spores in a suspension iii. To learn the steps in determining the concentration of bacteria in a suspension and how to calibrate the

concentration for a given bacteria using a spectrophotometer. 2. Requirements Counting chamber (haemocytometer), fungal cultures, sterile distilled water, sterile pipettes, sterile distilled water in universal bottles (9 ml in each bottle), sterile Petri dishes, nutrient agar medium (molten in water baths, colony counter 3. Counting the number of spores in a suspension using haemocytometer Work on haemocytometers will be in form of demonstrations. There are a number of microscopes provided and haemocytometers are mounted showing different number of cells. Each student will be expected to make counts of the cells and calculate the number of cells per ml. Note: Haemacytometers are expensive and must be handled with a lot of care. In case of problems, ask for assistance.

Haemocytometer

The circled grids indicate the large square (with 16 small squres) which should be used for counting

Procedure.

i. Flood about 10 ml of sterile distilled water on a culture containing good growth of a fungus that sporulates well

ii. Scrap the mycelia growth with a sterile glass slide or L-shaped glass rod to dislodge the spores iii. Sieve the suspension through a double layer of cheese cloth into a sterile beaker; if the suspension is

too turbid, add an equal volume of sterile distilled water. iv. Mix the suspension and draw a sample with a dropper; apply a small drop on the surface of the

haemocytometer – ensure the drop is placed at the centre on the area with grids. v. Apply a cover and blot any overflowing fluid from the edges of grooves of the counting chamber. vi. Place the haemocytometer on microscope stage and focus on the grid lines of the haemocytometer

using the 10X objective; focus on the large squares (1 large square has 16 small squares)


vii. Count the number of spores in one large square; repeat this for 3 other large squares and calculate the average number of spores in the 4 squares.

viii. Calculate the spore concentration using the formula: (Average Number of spores x104) dilution factor

The haemocytometer is designed so that the number of cells in one set of 16 corner

squares is equivalent to the number of cells x 104 / ml. The dilution factor is the

number of times the spore suspension was diluted, for example, if the spore

suspension was too turbid and you added an equal volume of water, the dilution

factor is 2.

4. Determination of the concentration of bacterial cells in a sample Requirements Cultures of bacteria on nutrient agar (e.g. Xanthomonas campestris pv campestris), sterile didtilled water, universal bottles containing 9 ml sterile distilled water, sterile pippetes, sterile L-shaped glass rods, sterile petri dishes, labels or marker pen, colony counter, nutrient agar media cooled to 50 0C in water bath. Procedure:

i. Flood the culture of bacteria with sterile distilled water and scrap the surface with sterile L-shaped glass rod.

ii. Pass the suspension through cheese cloth to filter off any agar fragments – sieve into a sterile beaker. iii. Label each universal bottle containing the 9 ml sterile distilled water as 10-1, 10-2, 10-3, 10-4, 10-5 and 10-

6 . iv. Take 1ml of suspension and add into the universal bottle labeled 10-1; mix the contents by shaking. v. From the 10-1, transfer 1ml of the suspension to the bottle labelled 10-2 with a sterile and fresh pipette.

This dilutes the original suspension by 100 times (102). vi. From the 10-2 suspension, transfer 1ml of the suspension to the bottle labelled10-3 using a fresh sterile

pipette, thus diluting the original suspension by 10-3 vii. Similarly repeat this procedure for the bottles labeled 10-4, 10-5 and 10-6 till the original sample is diluted

1,000,000 (10-6) and for each step use a fresh sterile pipette. viii. For each of the dilutions 10-1, 10-2, 10-3, 10-4, 10-5 and 10-6, measure the absorbance by means of

spectrophometer (this step will be demonstrated to you); record the absorbance readings. ix. From each of the dilutions 10-1, 10-2, 10-3, 10-4, 10-5 and 10-6, take 1ml of the suspension and place each

of the 1 ml suspension onto sterile Petri dishes correspondingly labeled 10-1, 10-2, 10-3, 10-4, 10-5 and 10-

6. Two Petri dishes should be used for each dilution. x. Into each Petri dish, gently add about 15ml of the molten nutrient agar medium cooled to 450C and

gently swirl to mix the soil suspension throughout the medium (Do not shake vigorously as this results in formation of air bubbles on the media surface); prepare two plates for each dilituin.

xi. Allow the plates to solidify and incubate in an inverted position for 1 to 3 days at room temperature xii. Count the number of colonies on each plate and calculate the number of cell by multiplying the number

of colonies by the dilution factor, e.g. if average of 30 colonies are counted in dilution 106, the number of cells will be: 30 x 106 = 3 x107 cells/ml. The results are reported as 3 x107 Colony Forming Units/ml or 3 x107 CFU/ml


101 102 103 104 105

9ml

Bacterialsuspension

1ml1ml 1ml 1ml 1ml

.....

Plate 1ml of each suspension in molten agar medium

1ml

106

xiii. Transfer the results to a table as shown below:

Dilution

101 102 103 104 105 106

No. of colony forming units (CFU)

Absorbance

xiv. Plot a graph of absorbance (Y-axis) against number of colony forming units (X-axis)

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