KING SAUD UNIVERSITY/ COLLEGE OF SCIENCE

Botany and Microbiology

LAB SKILLS (240 MBIO)

Subject Week Hours

Biosafety (safety rules and general precautions in laboratory) 1 4

Sterilization and Disinfection skills 2 4

Preparation of biological solutions and Buffer 3 4

Preparation of microbial cultivation media 4 4

Microscopy and Centrifugation techniques 5 4

Isolation and purification of microorganisms from soil and

human body

6 4

Preparation of microbial slides, Gram staining 7 4

Midterm Exam 1 8 4

Isolation and purification of fungi from different plant parts 9 4

Fungal staining 10 4

Chromatography and Spectrophotometric techniques 11 4

Midterm Exam 2 12 4

Enzymatic activity for microbes 13 4

Final Practical Exam 14

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 1

Lab 1

Title of the experiment: Biosafety (safety rules and general precautions in the laboratory)

Brief introduction:

1. Learning the technical safety requirements for work in microbiological laboratories.

2. Avoiding the harmful effects of microorganisms, chemical and radiation exposure in

microbiological labs.

Materials, tools and equipment:

Lab coat, Hand gloves, Safety goggles, Masks, Absolute Alcohol, etc

Procedures:

1. Use your lab coat, when working in the laboratory to protect clothing from

contamination or accidental discoloration by staining solutions.

2. Clean your work area (laboratory bench) with a recommended disinfectant such as 5%

Lysol or 5% phenol before and after each laboratory period to avoid contamination.

3. Eating, drinking, and smoking are forbidden at all times in the laboratory.

4. Always keep the laboratory work area free from articles not actually in us and replace

all reagents, cultures, and glassware in their appropriate places.

5. Keep your hands away from your mouth and eyes while in the laboratory. Do not place

anything such as pencils, food, and fingers in your mouth.

6. Wash your hands thoroughly before and after each experiment, using disinfecting soap

if possible.

7. Wear your musk and gloves to avoid inhalation of harmful solvents and buffer solutions

as well as affecting your hands.

8. Avoid contamination of benches, floor, and wastebaskets.

9. Place all discarded cultures, infectious materials, used glass slides and contaminated

glassware into the provided receptacles. Do not let unneeded materials accumulate.

10. Place all used or contaminated pipettes in appropriate glass jar filled with any

disinfectant such as 5% phenol.

11. When infectious material is accidentally spilled, cover it immediately with a

disinfectant such as 5% Lysol or 5% phenol and notify your instructor at once.

12. Do not move with a loop or pipette containing infectious material through the

laboratory. Flame wire loops and needles before and immediately after transfer of

cultures.

13. Label all experimental material properly.

Results:

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Conclusion:

The above mentioned safety rules will be beneficial for the students to work in safe and clean

environment.

References

1. Charlotte Bailey and Vicki Barwick (2007). Laboratory Skills Training Handbook. May

2007

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 2

Lab 2

Title of experiment: Sterilization and Disinfection Skills

Brief introduction:

Study the sterilization techniques and learn, how and when we use the different heat based and

another sterilization tools.

Sterilization: is defined as, the complete destruction or elimination of all forms of viable

microorganisms from a material by a chemical or physical methods.

Material, tools and equipment

Autoclave, Hot air over, Filtration assembly, portable UV chamber, Petri dishes, Erlenmeyer

flask, Beakers, Nichrome Wire Loop, Alcohol.

Procedure

Definition and explanations of Sterilization techniques:

1-Heat:

It is considering the most common and reliable method of sterilization. This method can be

used for all materials that withstand heat. Sterilization by heat could be concluded as follow.

1.1. Dry Heat:

a. Direct flaming:

It is a simple and common method for effective sterilization, of materials that can be heated to

redness in direct flame. Flaming could be used in microbiology labs to sterilize inoculating

needles, loops, forceps tips, searing spatulas and straight-wires by exposure to Bunsen burner

until become red. Loops, forceps tips and other metal object could be dipped in 70% ethanol

before briefly passing over Bunsen burner flame to obtained maximum degree of sterilization.

Mouth of test tubes, flasks, glass slides and cover slips are also sterilized by passing through

the Bunsen burner flame for a few seconds to become free from germs.

b. Hot air ovens:

Dry heat can be used to sterilize glassware or other non-porous heat conductive materials. In

dry ovens, 1 to 2 hours at 160 to 170°C is standard setting for complete sterilization. Dry heat

takes much longer time than moist heat to be transferred to the organism, thus both the time

and the temperature must usually be increased. Dry heat has the advantage that it can be used

on powders and other heat-stable materials that are adversely affected by steam.

1.2. Moist Heat:

Steam under pressure (Autoclaving):

This technique could be used to sterilize microorganism’s culture media, aqueous solutions,

and contaminated items such as Petri dishes, tips, tubes, gloves and plastic vessels comprised

of polypropylene.

Water boils at 100°C at atmospheric temperature, but within an autoclave, steam pressure can

reach (15 psi, above atmospheric pressure) with this increased pressure (the water temperature

will be increased to 121°C which will sterilize media more efficiently. Generally, we can use

an autoclave or pressure cooker at 121°C; 15 psi for 15 minutes, this is the standard

recommended for most microbiological media. These conditions are adequate to quickly and

effectively kill all microbial forms even the hardiest spore formers.

2. Radiation:

There are two types of radiation could be employed to achieve microbe-free environment.

Because radiation does not generate heat, it is termed "cold sterilization". Rays are harmful to

skin and eyes. It doesn't penetrate glass, paper or plastic.

Ionizing rays but. Electromagnetic rays such as gamma rays produced by a linear accelerator

from a heated cathode are high-energy with good penetrative power. It is used to sterilize

disposable Petri dishes, gloves, plastic syringes, dressing packs, antibiotics, vitamins,

hormones and fabrics in few seconds.

Non-ionizing rays: UV rays of wavelength longer than the visible light are low energy with

poor penetrative power. Wavelengths of UV rays, ranged from 200-280 nm have a destruction

effect on microbial cells, with 260 nm being most effective. UV rays use in surface disinfection

to disinfect hospital wards, operation theatres, virus laboratories, corridors, etc.

3. Chemicals:

There are many chemicals that have a broad spectrum of antimicrobial activity and are fast

acting. Mercuric chloride, sodium hypochlorite, formalin, phenols, ethanol and isopropanol are

extensively used in microbiological labs for a variety of purposes such as swabbing a bench

before and after use, for the sterilisation of surfaces, and for the disposal of used instruments

such as Pasteur pipettes. Many laboratory disinfectants need to be prepared before they can be

used.

When using disinfectants, it is important to ensure that they are used at the correct concentration

and that they are left to work for the correct length of time.

4. Filtration:

Filtration could be done for liquids and gases that may chemically altered by heat exposure

such as antibiotic and certain of culture media components. These solutions could be sterilized

by passing them through filters of an appropriate pore size. While most solutions used in

molecular biology will be adequately sterilized with a 0.22-μ filter, those for tissue culture

should be use A 0.1-μ filter to remove mycoplasma from tissue-culture. There are much kind

of filters could be used for such purpose i.e. asbestos filters, membrane filters and sintered

glass.

Results:

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Conclusion

The student learned different sterilization methods.

References

1. Rutala, W. A.; Weber, D. J. (2004) "Disinfection and Sterilization in Health Care Facilities:

What Clinicians Need to Know". Clinical Infectious Diseases. 39 (5): 702–709.

2. Mushtaq, Maryam; Banks, Charles J.; Heaven, Sonia (2012). "Effectiveness of pressurised

carbon dioxide for inactivation of Escherichia coli isolated from sewage sludge". Water

Science and Technology. 65 (10): 1759–1764.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 3

Lab 3

Title of experiment: Preparation of biological solutions and Buffers

Brief introduction:

Learn preparation biological solution, calculations and preparation of buffer solution.

Material, tools and equipment:

Salt, Sugar, Buffer tablets of varying pH, water, pH meter

Procedure

Solution:

A liquid mixture in which the minor component (the solute) is uniformly distributed within

the major component (the solvent).

Components of Solution

Solvent

The component present in larger proportion is known as solvent.

Solute

The component present in smaller proportion is known as solute.

Solution = Solvent + Solute

Ex. Sugar solution = Sugar (solute) + Water (solvent)

Common salt solution = Salt (solute) + Water (solvent)

Making a saline water solution by dissolving table salt (NaCl) in water.

The salt is the solute and the water the solvent.

Characteristics:

o A solution is a homogeneous mixture of two or more substances.

o The particles of solute in a solution cannot be seen by the naked

eye.

o A solution is stable.

o The solute from a solution cannot be separated by filtration (or mechanically).

o It is composed of only one phase.

Solubility:

o The ability of one compound to dissolve in another compound is called solubility.

o When a liquid can completely dissolve in another liquid the two liquids are miscible.

o Two substances that can never mix to form a solution are called immiscible.

Calculations:

Grams per liter (g/L.) Simply the mass of a solute in grams dissolved in a given volume of

solution. Usually used for solutions made from solid solutes and liquid solvents, like our sugar

and vinegar example.

Sugar: 20 gms/lit

Percent composition. The number of parts (again, usually grams) of solute found in one

hundred parts of solution. The percent symbol % means "out of 100" so you can easily write

the fraction as a percentage.

Sugar: 10% means, 10 gms/100 ml

PPM = parts per million

PPM is a term used in chemistry to denote a very, very low concentration of a solution.

One gram in 1000 ml is 1000 ppm and one thousandth of a gram (0.001g) in 1000 ml is one

ppm.

One thousandth of a gram is one milligram (1 gm = 1000 mg)

and 1000 ml is one liter, (1 lit= 1000 ml)

so that 1 ppm = 1 mg per liter = mg/Liter.

Molal solution:

Molality (m) is defined as the number of moles of solute per kilogram of solvent.

molality = moles of solute/kilograms of solvent

Molar solution

Molarity (M) is defined as the number of moles of solute per liter of solution.

molarity = moles of solute/liters of solution

o One molar (1 M) solution means one mole of a substance (solute) per liter of solution.

o A mole means gram molecular weight or molecular weight of a substance in grams. So

the molecular weight of a chemical is also its molar weight.

o To calculate the molecular weight one needs to add the atomic weights of all the atoms

in the molecular formula unit.

For example the molecule of NaCl consist of sodium (Na) and Chlorine (Cl). Their respective

atomic weights are NA-23 and Cl-35, so the molecular weight is 23+35= 58. Thus 58 grams of

NaCl equals one mole of NaCl.

Normality

Normality (N) is defined as the number of mole equivalents per liter of solution:

normality = number of mole equivalents/1 L of solution

Like molarity, normality relates the amount of solute to the total volume of solution; however,

normality is specifically used for acids and bases. The mole equivalents of an acid or base are

calculated by determining the number of H+ or OH- ions per molecule:

N = n × M (where n is an integer)

For an acid solution, n is the number of H+ ions provided by a formula unit of acid.

Example: A 3 M H2SO4 solution is the same as a 6 N H2SO4 solution.

For a basic solution, n is the number of OH- ions provided by a formula unit of base.

Example: A 1 M Ca(OH)2 solution is the same as a 2 N Ca(OH)2 solution.

Examples:

If 40 grams of NaOH with an equivalent weight or mass of 40 is dissolved in one liter of a

solution, normality of the solution is one and the solution is called as ONE

36.5 grams of hydrochloric acid (HCl) is a 1 N (one normal) solution of HCl.

Note: The normality of a solution is NEVER less than its molarity!

Dilution Formula: N1V1 = N2V2

This equation applies to all dilution problems.

C1 (initial conc) x V1 (initial volume) = C2 (final conc) x V2 (final volume)

Example: What volume of 1 N solution must be used to give 4.00 liters of a 0.1 N solution?

N1 = 1 N, V1 = unknown, N2 = 0.1 N, V2 = 4 liters = 4000 mls

N1V1= N2V2

V1 = (C2 x V2) / C1

= (0.100 X 4000) / 1 = 400 / 1 = 400 ml.

Buffer solution:

Definition

A buffer solution is one which resists changes in pH when small quantities of an acid or an

alkali are added to it.

Simple buffering agents

Buffering

agent

Useful pH range

Citric acid 2.1–7.4

Acetic acid 3.8–5.8

KH2PO4 6.2–8.2

CHES 8.3–10.3

Borate 8.25–10.25

Common buffer compounds used in biology

Common

name

Structure Mol.

weight

TAPS

243.3

Bicine

163.2

Tris

121.14

Tricine

179.2

TAPSO

259.3

HEPES

238.3

TES

229.20

MOPS

209.3

PIPES

302.4

Cacodylate

138.0

MES

195.2

Buffers are mainly classified of two types:

(а) mixtures of weak acids with their salt with а strong base

(b) mixtures of weak bases with their salt with а strong acid.

А few examples are given below:

Н2СО3 / NаНСО3 (Bicarbonate buffer; carbonic acid and sodium bicarbonate)

СН3СООН / СН3СОО Na (Acetate buffer; acetic acid and sodium acetate)

Na2HPO4/ NaH2PO4 (Phosphate buffer)

Results

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Conclusion

The student prepared varying concentration of the solution and buffers, and learned the

calculations about solution in laboratory.

References

1. Charlotte Bailey and Vicki Barwick (2007). Laboratory Skills Training Handbook. May

2007

2. Scorpio, R. (2000). Fundamentals of Acids, Bases, Buffers & Their Application to

Biochemical Systems. ISBN 978-0-7872-7374-3.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 4

Lab 4

Title of experiment: Microscopy and Centrifugation techniques

Brief introduction:

1. Study the different kinds of microscopes and centrifuge.

2. Learn how to use this device correctly.

Material, tools and equipment

Microscope, glass slides, cover slips, Cedar wood oil, tissue papers, etc.

Centrifuge, rotors, Tubes, Eppendorf.

Procedure

Definitions and explanations (kinds of microscopes):

1. Optical Microscope:

The optical microscope has one or two lenses that work to enlarge and enhance images placed

between the lower-most lens and the light source. It was the first device ever created.

a. Simple Optical Microscope:

It was used by Anton Van Leeuwenhoek during the late-sixteen and early-seventeenth

centuries, around the time that the microscope was invented. It has one lens, the convex lens,

in the magnifying process.

b. Compound Optical Microscope:

It has two lenses, work to minimize both chromatic and spherical aberrations so that the view

is unobstructed and uncorrupted. The compound light microscope consists of three sets of

lenses: -

- Condenser contains lenses that collect and focuses the light to upward directing through

any object on the stage. The amount of light is controlled by shutter, or iris diaphragm,

which adjust the amount of light admitted.

- The objectives include three or four lenses that provide a magnified and inverted image

of the specimen

- The eyepiece adds further magnification

2. Stereoscopic (Dissecting) microscope:

It is two microscopes in one, and uses two separate optical shafts, which focus on the same

point from different angles to produces a three-dimensional visualization of the examined

sample. It provides slightly different viewing angles to the left and right eyes because using

two separate optical paths with two objectives and two eyepieces.

Stereo microscope is relatively low power compared with compound microscopes (below 100

X). It is often used to study the surfaces of solid specimens or to carry out close work such as

sorting, dissection, microsurgery, small circuit board manufacture or inspection, and the like.

3. Inverted Microscope:

This kind of microscope views objects from an inverted position than that of regular

microscopes. It used to the study cell cultures in liquid media.

4. Fluorescence microscopy:

Fluorescence microscope is widely used device in the life sciences and biology. It is a light

microscope used to study properties of organic or inorganic substances using the phenomena

of fluorescence and phosphorescence. Both of an excitation and emission filters used in the

fluorescent microscope.

5. Digital microscope:

Traditional optical, stereoscopic and inverted microscopes have been recently modified into

digital microscopes. These modified microscopes constructed from computer units attached to

camera devices. Computer software converts the images to be displayed on a high resolution

LCD monitor instead of direct viewing.

6. Electron Microscopes:

Electron microscopy employs electron waves running parallel to a magnetic field providing

higher resolution. Electron microscopy allows one to visualize objects that are as small as 1

nm. Electron microscopy is a high-cost technology use very expensive materials such as

osmium gold-palladium or carbon or platinum.

a. Scanning electron microscope (SEM): Scanning electron microscope used to visualize the surface of tissues, macromolecular

aggregates.

b. Transmission electron microscope (TEM):

Transmission electron microscope used to study the inner structure of objects (tissues,

cells, viruses).

Centrifugation

Centrifugation is a process which involves the application of the centrifugal force for the

sedimentation of heterogeneous mixtures with a centrifuge, and is used in industrial and

laboratory settings. This process is used to separate two miscible substances, but also to analyze

the hydrodynamic properties of macromolecules. More-dense components of the mixture

migrate away from the axis of the centrifuge, while less-dense components of the mixture

migrate towards the axis. Chemists and biologists may increase the effective gravitational force

on a test tube so as to more rapidly and completely cause the precipitate (pellet) to gather on

the bottom of the tube. The remaining solution (supernatant) may be discarded with a pipette.

There is a correlation between the size and density of a particle and the rate that the particle

separates from a heterogeneous mixture, when the only force applied is that of gravity. The

larger the size and the larger the density of the particles, the faster they separate from the

mixture. By applying a larger effective gravitational force to the mixture, like a centrifuge does,

the separation of the particles is accelerated. This is ideal in industrial and lab settings because

particles that would naturally separate over a long period of time can be separated in much less

time.

The rate of centrifugation is specified by the angular velocity usually expressed as revolutions

per minute (RPM), or acceleration expressed as g. The conversion factor between RPM and g

depends on the radius of the centrifuge rotor. The particles' settling velocity in centrifugation

is a function of their size and shape, centrifugal acceleration, the volume fraction of solids

present, the density difference between the particle and the liquid, and the viscosity. The most

common application is the separation of solid from highly concentrated suspensions, which is

used in the treatment of sewage sludges for dewatering where less consistent sediment is

produced.

Centrifugation

Centrifugation in biological research

Microcentrifuges

Microcentrifuges are used to process small volumes of biological molecules, cells, or nuclei.

Microcentrifuge tubes generally hold 0.5 - 2.0 mL of liquid, and are spun at maximum angular

speeds of 12,000–13,000 rpm. Microcentrifuges are small enough to fit on a table-top and have

rotors that can quickly change speeds. They may or may not have a refrigeration function.

High-speed centrifuges

High-speed or superspeed centrifuges can handle larger sample volumes, from a few tens of

millilitres to several litres. Additionally, larger centrifuges can also reach higher angular

velocities (around 30,000 rpm). The rotors may come with different adapters to hold various

sizes of test tubes, bottles, or microtiter plates.

Rotor

Results

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Conclusion

The student learned the microscopy and centrifugation techniques alongwith on hand training

session.

References

1. Van Helden, Albert; Dupre, Sven; Van Gent, Rob (2011). The Origins of the Telescope.

Amsterdam University Press. ISBN 978-9069846156.

2. Susan R. Mikkelsen & Eduardo Cortón. Bioanalytical Chemistry, Ch. 13. Centrifugation

Methods. John Wiley & Sons, Mar 4, 2004, pp. 247–267.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 5

Lab 5

Title of the experiment: Preparation of microbial cultivation media

Brief introduction:

1. Study the different kinds of media.

2. Learn how to use them during isolation and purification of microorganisms.

Material, tools and equipment:

Nutrient agar, Nutrient broth, Potato Destrose Agar, Mac Conkey Agar, Agar agar, Distilled

water. etc,

Procedure:

Medium: a nutrient blend used to support microbial growth.

A medium is sterilized (living organisms removed) before usage in the lab.

Important points:

Sterilization methods include; autoclaving, dry-heat, filtration, UV exposure and ethylene

oxide.

Culture: Is part of specimen grown in culture media.

Culture Media: is a medium (liquid or solid) that contains nutrients to grow bacteria in vitro.

Because sometimes we cannot identify with microscopical examination directly, and

sometimes we do culture for antibiotic sensitivity testing.

History o The original media used by Louis Pasteur – meat broth

o Liquid medium – diffuse (spread out over a large area) growth

o Solid medium – discrete (individually separate and distinct) colonies.

o Cooked cut potato by Robert Koch – earliest solid medium

o Gelatin – not satisfactory

o -liquefy at 24oC

Properties of agar:

o Angelina Fanny Hesse

o Used for preparing solid medium

o Obtained from seaweeds

o Comes as sold powder and then you add water to it.

o No nutritive value

o Not affected by the growth of the bacteria.

o Melts at 98oC & sets at 42oC

o 2% agar is employed in solid medium

Properties of Media

o Support the growth of the bacteria.

o Should be nutritive (contains the required amount of nutrients).

o Suitable pH (neutral to slightly alkaline 7.3-7.4).

o Suitable temperature, and suitable atmosphere. (Bacteria grow at 370C)

o Note: media are sterilized by autoclaving at 1210C and 2 atmosphere for 15-20 minutes.

With the autoclave, all bacteria, fungi, viruses, and spores are destroyed. Some media

can’t be sterilized by autoclaving because they contain eggs or carbohydrates.

Types of culture media

I. Based on their consistency

a) solid medium

b) liquid medium

c) semi solid medium

II. Based on the constituents/ ingredients

a) simple medium

b) complex medium

c) synthetic or defined medium

d) Special media

Special media

• Enriched media

• Enrichment media

• Selective media

• Indicator media

• Differential media

• Sugar media

• Transport media

• Media for biochemical reactions

III. Based on Oxygen requirement

- Aerobic media

- Anaerobic media

Results

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Conclusion

The students learned to prepare various kinds of media for the growth of microorganisms.

References

1. Madigan M, Martinko J, eds. (2005). Brock Biology of Microorganisms (11th ed.). Prentice

Hall. ISBN 0-13-144329-1.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 6

Lab 6

Title of experiment: Isolation, purification of microorganisms from soil and human body

Brief introduction

Isolation of fungi from soil from soil:

Learning how to isolate soil fungi.

Materials, tools and equipment:

Soil sample.

Acidified Czapek–Dox + 0.5 % yeast extract agar medium.

Sterilized Petri plates.

Sterilized distilled water in test tubes (9cm water/each).

Golf shaped like glass road.

Procedures

Simple plating technique (Direct isolation): The direct inoculation method may be best for

isolating various and general soil fungi simply, readily, and economically.

1- Transfer a small amount (0.005–0.015 gm) of soil to a sterilized Petri dish.

2- Added 8–10 ml. of semi-cooled (45°C) nutrient medium and shake the plate to let the

soil particles dispersed throughout the thin layer of agar medium before it solidifies.

3- If the soil is very dry, or contains a high proportion of clay, it is preferable to mix the

particles with a drop of sterile water in the plate, before adding the medium.

4- Incubate treated plates at 20-30°C, investigate the colonies appearance after 48 and

record the results.

-Dilution (Plate) Method: -

- Take a proper amount of air dried soil sample after saving it to remove any undesirable

materials (plant duperies and beg granuls).

- Prepare cereal dilution (i.e. 1:10, 1:100, 1:1000……. etc) from the soil sample.

-Transfer one drop from each of the last two dilution samples to plated isolation media using a

sterile pipette.

- Use golf shaped like glass road to spread the droplets onto the agar surface.

- Incubate treated plates at 20-30°C, investigate the colonies appearance after 48 and record the

results.

Isolation of soil bacteria

Objective:

Learning how to isolate soil bacteria.

Materials:

Soil samples.

Water agar medium.

Sterilized Petri plates.

Bacterial loop.

Sterilized distilled water in test tubes (9cm water/each).

Procedure:

Isolation of soil bacteria using dilution technique.

1- Suspend 10 grams of air-dried powdered soil in 100 ml sterilized distilled water and shaken

for 15 minutes.

2- Make cereal dilutions of previous stock in glass test tubes.

3- Take one loop of the soil dilution to inoculate agar plates.

4- Use the latest two dilutions.

5- Incubate the inoculated plates at 30°C for 24 hours.

6- Investigate your plates and write a detailed report.

Pure culture technique

A pure culture is the basis for diagnosis, identification and all further work of microbiological

studies. For pure culture, all works should be performed under aseptic conditions (in a flow

cabinet, if possible) to prevent any contaminating microorganism to reach the medium.

1 2 3 4

Isolation of Fungi:

Objective:

Learning the purification techniques of fungal isolates.

Materials:

Fresh fungal culture.

Water agar medium.

Sterilized Petri plates.

Bacterial loop.

Sterilized distilled water in test tubes (9cm water/each).

Procedures:

There are two different methods to obtain fungal pure culture from the wild isolates.

Single spore culture: This method used in sporulating fungi such as Fusarium, Cladosporium

and Cercospora as a following:

1. Cereal dilution of spore suspension is prepared from previously plated fungal isolate using

sterile distilled water.

2. Solidified water agar plates are inoculated by droplets from the last two dilutions.

3. Golf stick like glass road is used to spread the spore suspension droplet onto the agar

surface. Bacterial loop could be also used to streak spore suspension onto the agar surface.

4. Inoculated plates are investigated under light microscope to locate single spore.

5. Marked single spore is transferred to the surface of new agar plate and incubated at 25-

30°C for 12-24h to germinate and start to grown into new single colony.

6. Obtained single spore isolates are immediately slanted and preserved.

Hypal tip culture:

This method used in unsporulating fungi or that producing very few spores.

- By the aide of dissecting microscope, most freely grown hyphal tip are cut from the

margin of previously plated fungal isolate and transfer to new water agar plates.

- Inoculated plates are incubated at 25-30°C for 24-72h until reaching a new single

colony.

- Obtained colonies are immediately slanted and preserved as pure cultures.

Results

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Conclusion

The student learned the isolation techniques from the soil samples, that brought to the laboratory

by different student from various places around campus.

References

1. Stefanis C, Alexopoulos A, Voidarou C, Vavias S, Bezirtzoglou E. (2013). Principal methods

for isolation and identification of soil microbial communities Folia Microbiol (Praha).

58(1):61-8.

2. Warcup J.H (1950). The Soil-Plate Method for Isolation of Fungi from Soil. Nature. 166: 117-

118.

Bacterial isolates:

Objective:

Learning the purification techniques of bacterial isolates.

Materials:

Fresh bacterial culture.

Water agar medium.

Sterilized Petri plates.

Bacterial loop.

Selective media.

Sterilized distilled water in test tubes (9cm water/each).

Procedure:

Bacterial pure culture could be obtained from mixed culture by using:

- Single colony selection.

- A selective medium such as mannitol salt agar is used to select Staphylococcus as a

selective medium, MacConkey agar is a selective medium for E. coli

- A differential medium is designed to show visible differences between micro-

organisms such as different bacteria producing different colours of colonies.

Single colony selection:

1. To obtained pure culture isolate, one separate colony must choose from the latest

streaked plate of such bacterial isolate.

2. Separate chosen colony is transferred to a new nutrient agar plate and streaked by

sterilized loop.

3. One loopful is used for each plate and streaking must be in one-way direction. Streaked

plates are incubated at 37°C for 12-24h or until colony appear. If this colony was

developed from one cell, only one type of colony will develop few days after streaking

onto the nutrient agar surface.

4. If only one type of colony was developed onto the nutrient agar plate, pure culture was

obtained and must transfer to a fresh nutrient slant.

Diagram explain how can pure bacterial culture obtain

Results

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Conclusion

The student learned the isolation techniques from the soil samples, that brought to the laboratory

by different student from various places around campus.

References

3. Stefanis C, Alexopoulos A, Voidarou C, Vavias S, Bezirtzoglou E. (2013). Principal methods

for isolation and identification of soil microbial communities Folia Microbiol (Praha).

58(1):61-8.

4. Warcup J.H (1950). The Soil-Plate Method for Isolation of Fungi from Soil. Nature. 166: 117-

118.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 8

Lab 8

Title of experiment: Preparing of microbial slides, Gram staining

Brief introduction:

Microscope examinations of microorganisms require some preparation to get a clear and

informative vision. Prepare of stained slide is one of the most common tool in this particular.

Bacterial staining:

Simple Stains

Bacterial shapes and activities could be observed under a cover glass without staining, but

forming a complete idea about their morphology is so difficult. Bacterial cells are minute and

tend to be transparent, even when magnified. So the simplest method for examining them is to

make stain preparations for microscopic study.

Objective:

To learn the simple stain technique and its value in studying basic microbial morphology.

Materials:

Alcohol solution.

Glass slides and cover slips.

Bacterial loop.

Sterilized distilled water.

Staining rack

Methylene blue.

Safranin.

Compound microscope

Procedures:

1. Wash appropriate numbers of glass slides by dipping in alcohol and polishing with

tissue or soft cloth.

2. Take three clean slides, label them and make a circle (about 1 cm in diameter) in the

center of each one using a fine marker.

3. Turn the slides over so that the unmarked side is up and with your inoculating loop,

place a loopful of water in the ringed area of the slide.

Simple staining

4. Take loopful of bacterial growth and mixing it (using proper aseptic transfer

techniques) in the water and spread it out. Repeat this step until make a smear of

bacteria.

5. Allow the smears to air dry until you see a thin white film on each slide. If not, add

another loopful of water and more bacteria as in step.

6. Heat-fix the smears by passing the slides rapidly through the Bunsen flame three times

so that the smears will not wash off. (Fixing could be done by flooding the slides with

absolute alcohol on a staining rack for one minute, then alcohol is drained off and the

slides air dries completely).

7. Place the slides on a staining rack and flood them with methylene blue. Leave the stain

on for three minutes.

8. Wash each slide gently with distilled water, drain off excess water, place the slides on

bibulous paper, and let them dry completely in air.

9. Prepare another set of slides as described above (steps 2-8) but stain them with safranin

for three minutes instead of methylene blue.

10. Wash, drain, and dry the slides as in step 8.

11. Examine all slides, microscopically and record the results in lab report.

Gram Stains

Bacteria morphology could be determined using the simple staining, but further staining

method is required to distinguish between bacteria of similar morphology. Gram stain

(differential staining) could be used in this particular. It stains gram positive bacteria by violet

color wile gram negative by red color.

Objective:

To learn the gram stain technique and its value in distinguishing bacterial groups.

Materials:

Alcohol solutions (70-95%).

Glass slides and cover slips.

Bacterial loop.

Sterilized distilled water.

Staining rack

Crystal violet.

Gram’s iodine

Safranin.

Bibulous paper

Compound microscope

Procedures:

1. Take for clean slides, mark them and make a circle (about 1 cm in diameter) in the

center of each one using a fine marker.

2. Turn the slides over so that the unmarked side is up and with your inoculating loop,

place a loopful of water in the ringed area of the slide.

3. Take loopful of bacterial growth and mixing it (using proper aseptic transfer

techniques) in the water and spread it out. Repeat this step until make a smear of

bacteria.

4. Allow the smears to air dry until you see a thin white film on each slide. If not, add

another loopful of water and more bacteria as in step.

5. Heat-fix the smears by passing the slides rapidly through the Bunsen flame three times

so that the smears will not wash off. (Fixing could be done by flooding the slides with

absolute alcohol on a staining rack for one minute, then alcohol is drained off and the

slides air dries completely).

6. Flood the slides with crystal violet (Leave for one minute) then wash off with tap water.

7. Flood with Gram’s iodine (Leave for one minute) then wash off with tap water

8. Wash with alcohol (95% for 10–20 seconds) until no more color washes off (avoid

over washing) then wash off with tap water.

9. Apply safranin (for one minute) then wash off with tap water.

10. Drain off excess water, place the slides on bibulous paper, let them dry completely in

air (just before you examine the preparation microscopically). Label the slides when

become dry.

11. Examine all slides under oil with the oil-immersion objective and record your

observations in lab report.

Results

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Conclusion

The students learned the staining technique for bacteria. Gram positive and negative bacteria

retain the primary and secondary stain due to the respective cell wall architecture

(Peptidoglycan layer in cell wall).

References

1. Madigan M, Martinko J, eds. (2005). Brock Biology of Microorganisms (11th ed.). Prentice

Hall. ISBN 0-13-144329-1.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 9

Lab 9

Title of experiment: Isolation of fungi from diseased plant parts

Brief introduction:

Learning how to isolate plant pathogenic fungi from diseased plant parts.

Material, tools and equipment:

Diseased plant sample.

Sterilized filter papers.

Sodium hypochlorite solution 5%.

Water agar medium.

Sterilized Petri plates.

Sterilized distilled water.

Procedures:

The general procedures for isolation of any fungi from diseased plant materials are as follows:

1. Use running tap water to wash plant materials for at least 30 min.

2. Select the most freshly infected plant parts (from diseased plants), sterilize the selected parts

using 5% sodium hypochlorite solution or 70% ethanol for 30 sec to 3 min.

3. Cut each surface sterilized plant part into tissue segments of less than 5 mm.

4. Transfer prepared tissue segments to plated isolation media using sterilized forceps (3-4

segments/9cm. plate).

5. Incubate cultured plates at the appropriate temperature (25-30°C) for 1 to 7 days.

6. Investigate the incubated plates regularly because fungal hyphae may be elongated from the

plated tissue segments within a few days.

7. Transfer any appeared hyphal tip to a new agar plate as soon as possible to avoid any

contamination.

Results

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Conclusion

The students learned the isolation technique for the fungi, separate them individually.

Reference

1. Hawksworth DL, Lücking R (2017). Fungal Diversity Revisited: 2.2 to 3.8 Million

Species.. Microbiol Spectr. Microbiol Spectrum 5(4):FUNK-0052-2016.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 10

Lab 10

Title of experiment: Fungal staining

Brief introduction:

To learn the simplest preparation technique of stained fungal slides and its value in fungal

identification.

Material, tools and equipments:

Fungal cultures on Sabouraud dextrose agar (Aspergillus, Penicillium, Rhizopus &

Saccharomyces).

Clear tape.

Glass slides.

Lactophenol cotton blue (for staining molds).

Light microscope.

Miscellaneous supplies.

Lens paper.

Procedure:

1. Examine the plated cultures of the four fungi, recording all colony characters that may help

recognizing fungal genes.

2. After examining the colonies, make a pressure tape preparation of the 4 fungal cultures as

following:

a. Place a drop of lactophenol cotton blue on the center of the slide using a fine dropper.

b. Hold a piece of clear sticky tape in a U-shape, sticky side down as illustrated in the fig.

c. Touch the surface of a fungal colony carefully by the sticky tape.

d. Place the tape sticky side down in a lactophenol cotton blue drop.

e. Fold extra length of tape around edges of slide.

f. Examine this preparation using the light microscope then record your observations.

Results

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Conclusion

The student learned the staining technique for the fungi. Each student performed staining

individually by taking precautionary measures.

References

1. Cumitech 11: Practical Methods for Culture and Identification of Fungi in the Clinical

Microbiology Laboratory. 1980. American Society for Microbiology, Washington,

D.C.

2. Kwon-Chung, K.J. and J.E. Bennett. 1992. Medical Mycology. Lea and Febiger,

Malvern, PA.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 11

Lab 11

Title of experiment: Chromatography Technique

Brief introduction:

1. Study the different kinds of chromatography and spectroscophometric procedure.

2. Learn how to use this device correctly.

A diagram to illustrate the principles of Paper chromatography

(a) At the start mark a baseline in pencil - mark won't dissolve or run. Make sure it is above

the surface of the solvent, so as not to dissolve the spots directly. You then carefully put spots

of the mixtures and standards (known materials) onto the baseline. so we start with an even

line of solute chemical spots e.g. dyes or any coloured compound.

(b) The prepared chromatogram paper is carefully placed in the solvent so the baseline and

spots are above the solvent. The solvent is allowed to move up the paper. The colours begin to

separate out. Its best done in a larger covered beaker so the solvent doesn't evaporate into the

laboratory!

(c) When the solvent is near the top of the paper, the paper is removed and allowed to dry

before examination and measurement of the Rf values of all the dyes. The final result is called

the chromatogram.

(d) How do we measure an Rf value? Its quite simple to measure a reference value (Rf) value

for a dye colour. e.g. for the green dye spot, S might be 7.2 cm, D might be 5.3 cm, so

Rf = D/S = 5.3/7.2 = 0.74

For best results:

(i) try different solvents eg water or ethanol ('alcohol'), propanone ('acetone') or butanol.

(ii) If you can enclose the whole system in a larger glass container with a lid on, it reduces

evaporation of the solvent.

(iii) The fastest moving spots should be able to move at least 5 cm to get a reasonably accurate

Rf value.

For accurate work the distance moved by the solvent is marked on carefully with a pencil and

the distances moved by each 'centre' of the coloured spots is also measured. These can be

compared with known substances BUT if so, the identical paper and solvent must be used (See

Rf values below and diagrams above). The Rf values is the ratio how far the spot travels (D on

diagram) relative to the distance moved by the solvent (S on the diagram) and must have values

of >0 and <1

Reference values Rf = distance move by substance

Chromatography Rf vales depend on the substance, the type of paper (stationary phase) AND

the solvent (mobile phase). To get consistent chromatography results you must use identical

conditions, including temperature, and identical chemical reagents. These criteria is an example

of 'fair testing' methodology. You may have to experiment with solvents to ensure the best

separation of the components.

Material, tools and equipment:

TLC plates, Green leaves, Alcohol, Mortar and Pestle, etc.

Procedure:

A plant material extraction process using chromatography The experiment described below simulates one way in which drug companies extract plant

material to develop products in the pharmaceutical industry.

1. Take some suitable plant material and crush it with a pestle and mortar.

2. Scrape the crushed plant material and gently heat with a small volume of suitable solvent

like water or alcohol to dissolve and extract some of the soluble coloured plant material.

3. Filter off the residue to give a clear but coloured concentrated solution.

4. Take the solution and spread it along the start line of some chromatography paper (diagram

A).

5. Dip and suspend the chromatography paper into a suitable solvent in a covered container so

the start line is above the solvent surface and let the solvent be absorbed by the chromatography

paper and move upwards (diagram B).

6. When the solvent front has reached near the top of the paper, stop (diagram C) and hang up

the chromatogram to dry.

7. You can then separate the mixture 'physically' by cutting strips off for each band of coloured

material that has separated out on the chromatogram (diagram D).

8. You can then extract each separated product by re-dissolving it by placing the strips in the

solvent.

9. The different solutions can then be carefully evaporated to produce the separated coloured

solid materials.

Results

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Conclusion

The students learned the chromatographic technique by doing individual experiments by leaf

extract and separated the different leaf compound in different solvent system.

References

1. McMurry, John (2011). Organic chemistry: with biological applications (2nd ed.).

Belmont, CA: Brooks/Cole. p. 395. ISBN 9780495391470.

2. Ettre, L. S.; Sakodynskii, K. I. (1993). "M. S. Tswett and the discovery of

chromatography II: Completion of the development of chromatography (1903–1910)".

Chromatographia. 35 (5–6): 329–338.

Symbol and number of course: MBIO 240

Name of course: Laboratory Skills

Number of experiment: 13

Lab 13

Title of experiment: Microbial enzymatic activities

Brief introduction

Some microorganisms capable of hydrolyze large organic molecules due to enzymatic activity

and then use the component parts in further metabolic processes.

Amylase activity

Starch is a polysaccharide that is hydrolyzed by amylase producing microorganisms. When

iodine is added to the intact starch molecule, a blue-colored complex forms. If starch is

hydrolyzed into simple sugars i.e. glucose and maltose, however, no color reaction is seen.

Materials, tool and equipment:

Starch agar plates - Fresh culture of Aspergillius niger. - Gram’s iodine solution

Procedures:

1. Prepare starch agar plates. Invert each one, and divided it into two equal sections with

marking pencil.

2. Inoculate one section with 6mm plug of Aspergillius niger and another section with water

agar plug as a control.

3. Label each section of the plate correctly and incubate treated plates for three days at 28°C.

5. When the cultures have grown, drop Gram’s iodine solution onto the plate until the entire

surface is lightly covered.

6. Record your investigations in lab report.

Results

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Conclusion

The student learned the enzyme assay by employing the fungus Aspergillus niger in the starch

agar plate. The starch hydrolysis took place near the growth of the fungal mycelial mat, hence

the clear zone appears by addition of iodine.

References:

1. Tille P.M. 2014. Bailey and Scott’s diagnostic microbiology. Thirteen edition. Mosby, Inc.,

an affiliate of Elsevier Inc. 3251 Riverport Lane. St. Louis. Missouri 63043

Urease activity: -

Some bacteria produce urease enzyme and split urea molecule, releasing carbon dioxide and

ammonia. When bacterial cells that produce urease are grown in this medium, urea is degraded,

ammonia is released, and the pH becomes alkaline. This pH shift is detected by a change in the

Phenol red color from orange-pink to dark pink.

Materials:

Urea broth or urea agar medium and Proteus vulgaris culture.

Procedures

1. Inoculate a tube of urea broth or agar with the Proteus culture. Include un-inoculated

treatment.

2. Incubate the tubes at 35°C for 24 hours.

5. When the cultures have grown, drop Phenol red solution in the broth or onto the plate until

the entire surface is lightly covered.

6. Record your investigations in lab report.

Results

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Conclusion

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References:

1. Tille P.M. 2014. Bailey and Scott’s diagnostic microbiology. Thirteen edition. Mosby, Inc.,

an affiliate of Elsevier Inc. 3251 Riverport Lane. St. Louis. Missouri 63043