BI0217-BASIC BIOTECHNOLOGY LAB MANUAL

Offered to II YEAR B.TECH BIOINFORMATICS

DEPARTMENT OF BIOINFORMATICS SCHOOL OF BIOENGINEERING

SRM UNIVERSITY KATTANGULATHUR

2

S.NO

NAME OF THE EXPERIMENT PAGE NO

DATE OF EXPERIMENT

SIGN

1 LABORATORY SAFETY-GENERAL RULES AND REGULATIONS LABORATORY PROTOCOL

2 STERILIZATION TECHNIQUES 3 PREPARATION OF MEDIA

4 ISOLATION, ENUMERATION AND PURIFICATION OF MICROBES FROM A GIVEN SAMPLE

5 USE OF MICROSCOPE 6 SIMPLE STAINING 7 GRAM STAINING

8 SPORE STAINING (SCHAEFFER-FULTON METHOD)

9 HANGING DROUP

10 PREPARATION OF BUFFERS AND MEASUREMENT OF pH

11 ESTIMATION OF SUGARS

12 ESTIMATION OF PROTEINS BY LOWRY’S METHOD / BIURET METHOD

13 ESTIMATION OF CHOLESTEROL BY ZAK’S METHOD

14 SEPARATION OF AMINO ACIDS - THIN LAYER CHROMATOGRAPHY

15 SEPARATION OF SUGARS - PAPER CHROMATOGRAPHY

16 BIOCHEMICAL ESTIMATION OF DNA /RNA USING SPECTROPHOTOMETER

INSTRUCTIONS: ALL DIAGRAMS IN THE LEFT HAND SIDE OF THE PAGE

3

LABORATORY SAFETY - GENERAL RULES AND REGULATIONS A rewarding laboratory experience demands strict adherence to prescribed rules for personal and environmental safety. The former reflects concern for your personal safety in terms of avoiding laboratory setting to prevent contamination of experimental procedures by microorganisms from exogenous sources. Because most microbiological laboratory procedures require the use of living organisms, an integral part of all laboratory session is the use of aseptic techniques. Although the virulence of microorganisms used in the academic laboratory environment has been greatly diminished because of their long-term maintenance on artificial media, all microorganisms should be treated as potential pathogens (organisms capable of producing disease). Thus, microbiology students must develop aseptic techniques (free of pathogenic organisms) in preparation for industrial and clinical marketplaces where manipulation of infectious organisms may be the norm rather than the exception. The following basic steps should be observed at all times to reduce the ever-present microbial flora of the laboratory environment.

1. Upon entering the laboratory, place coast, books, and other paraphernalia in specified locations-never on bench tops.

2. Keep doors and windows closed during the laboratory session to prevent contamination from air currents.

3. At the beginning and termination of each laboratory session, wipe bench tops with a disinfectant solution provided by the instructor.

4. Do not place contaminated instruments, such as inoculating loops, needles, and pipettes, on bench tops. Loops and needles should be sterilized by incineration, and pipettes should be disposed of in designated receptacles.

5. On completion of the laboratory session, place all cultures and materials in the disposal area as designated by the instructor.

6. Rapid and efficient manipulation of fungal cultures and materials in the disposal area as designated by the instructor.

7. Rapid and efficient manipulation of fungal cultures is required to prevent the dissemination of their reproductive spores in the laboratory environment.

To prevent accidental injury and infection of yourself and others, observe the following regulations at all times:

1. Wash your hands with liquid detergent and dry them with paper towels upon entering and prior to leaving the laboratory.

2. Wear a paper cap or tie back long hair to minimize its exposure to open flames 3. Wear a lab coat or apron while working in the laboratory to protect clothing from

contamination or accidental discoloration by staining solutions. 4. Closed shoes should be worn at all times in the laboratory setting. 5. Never apply cosmetics or insert contact lenses in the laboratory. 6. Do not smoke, eat, or drink in the laboratory. These activities are absolutely

prohibited. 7. Carry cultures in a test - tube rack when moving around the laboratory. Likewise,

keep cultures in a test-tube rack on the bench tops when not in use. This serves a dual purpose to prevent accidents and to avoid contamination of yourself and the environment.

8. Never remove media, equipment, or especially, bacterial cultures from the laboratory. Doing so is absolutely prohibited.

4

9. Immediately cover spilled cultures or broken cultures tubes with paper towels and then saturate them with disinfectant solution. After 15 minutes of reaction time, remove the towels and dispose of them in a manner indicated by the instructor.

10. Report accidental cuts or burns to the instructor immediately. 11. Never pipette by mouth any broth cultures or chemical reagents. Doing so is strictly

prohibited. Pipetting is to be carried out with the aid of a mechanical pipetting device.

12. Do not lick labels. Use only self-stick labels for the identification of experimental cultures.

13. Speak quietly and avoid unnecessary movement around the laboratory to prevent distractions that may cause accidents.

The specific precautions outlined below must be observed when handling body fluids of unknown origin due to the possible imminent transmission of the HIV and hepatitis B viruses in these test specimens.

1. Disposal gloves must be worn during the manipulation of these test materials. 2. Immediate hand washing is required if contact with any of these fluids occurs and

also upon removal of the gloves. 3. Masks, safety goggles, and laboratory coast should be worn if an aerosol might be

formed or splattering of these fluids is likely to occur. 4. Spilled body fluids should be decontaminated with a 1:10 dilution of household

bleach, covered with paper toweling, and allowed to react for 10 minutes before removal.

5. Test specimens and supplies in contact with these fluids must be placed into a container of disinfectant prior to autoclaving.

I have read the above laboratory safety rules and regulations and agree to abide by them.

5

LABORATORY PROTOCOL

Student preparation for laboratory sessions The efficient performance of laboratory exercises mandates that you attend each session fully prepared to execute the required procedures. Read the assigned experimental protocols to effectively plan and organize the related activities. This will allow you to maximize use of laboratory time. PREPARATION OF EXPERIMENTAL MATERIALS Microscope Slides: Meticulously clean slides are essential for microscopic work. Commercially precleaned slides should be used for each microscopic slide preparation. However, wipe these slides with dry lens paper to remove dust and finger marks prior to their use. Labeling of culture vessels: Generally, microbiological experiments require the use of a number of different test organisms and a variety of culture media. To ensure the successful completion of experiments, organize all experimental cultures and sterile media at the start of each experiment. Label culture vessels with non- water- soluble glassware markers and / or self - stick labels prior to their inoculation. The labeling on each of the experimental vessels should include the name of the test organisms, the name of the medium, the dilution of sample, if any, your name or initials, and the date. Place labeling directly below the cap of the culture tube. When labeling Petri dish cultures, only the name of the organism(s) should be written on the bottom of the plate, close to its periphery, to prevent obscuring observation of the results. The additional information for the identification of the culture should be written on the cover of the Petri dish. INOCULATION PROCEDURES Aseptic techniques for the transfer or isolation of microorganisms, using the necessary transfer instruments is described fully in the experiments in Part I of the manual. Technical skill will be acquired through repetitive practice. Inoculating Loops and Needles: It is imperative that you incinerate the entire wire to ensure absolute sterilization. The shaft should also be briefly passed through the flame to remove any dust or possible contaminants. To avoid killing the cells and splattering the culture, cool the inoculating wire by tapping the inner surface of the culture tube or the Petri dish cover prior to obtaining the inoculum. When performing an aseptic transfer of microorganisms, a minute amount of inoculum is required. If an agar culture is used, touch only a single area of growth with the inoculating wire to obtain the inoculum. Never drag the loop or needle over the entire surface, and take care not to dig into the solid medium. If a broth medium is used, first tap the bottom of the tube against the palm of your hand to suspend the microorganisms. Caution: Do not tap the culture vigorously as this may cause spills or excessive foaming of the culture, Which may denature the proteins in the medium. Pipettes: Use only sterile, disposable pipettes or glass pipettes sterilized in a canister. The practice of pipetting by mouth has been discontinued to eliminate the possibility of auto-infection by the accidental imbibement of the culture or infectious body fluids. Instead, a mechanical pipetting device is to be used to obtain and deliver the material to be inoculated. INCUBATION PROCEDURE Microorganisms exhibit a wide temperature range for growth. However for most used in this manual, optimum growth occurs at 370C over a period of 18 to 24 hours. Unless

6

otherwise indicated in specific exercise, incubate all cultures under the conditions cited above. Place culture tubes in a rack for incubation. Petri dishes may be stacked; however, they must always be incubated in an inverted position (top down) to prevent water of condensation from dropping onto the surface of the culture medium. This resultant excess moisture may then serve as a vehicle for the spread of the micro-organisms on the surface of the culture medium, thereby producing confluent rather than discrete microbial growth. PROCEDURE FOR RECORDING OBSERVATIONS AND RESULTS The accurate accumulation of experimental data is essential for the critical interpretation of the observations upon which the final results will be based. To achieve this end, it is imperative that you complete all the preparatory readings that are necessary for your understanding of the basic principles underlying each experiment. Meticulously record all the observed data in the "Observations and Results" section of each experiment. In the exercises that require drawings to illustrate microbial morphology, it will be advantageous to depict shapes, arrangements, and cellular structures enlarged to 5 to 10 times their actual microscopic size, as illustrated below. For this purpose a number 2 pencil is preferable. Stippling may be used to depict different aspects of cell structure, e.g., endospores or differences in staining density. REVIEW QUESTIONS The review questions are designed to evaluate understanding of the principles and the interpretations of observations in each experiment. Completion of these questions will also serve to reinforce many of the concepts that are discussed in the lectures. The designated critical-thinking questions are designed to stimulate further refinement of cognitive skills. PROCEDURES FOR TERMINATION OF LABORATORY SESSION 1. All equipment, supplies and chemical reagents are to be returned to their original locations. 2. All capped test-tube cultures and closed Petri dishes are to be neatly placed in a designated collection area in the laboratory for subsequent autoclaving. 3. Contaminated materials, such as swabs, disposable pipettes, and paper towels, are to be placed in a biohazard receptacle prior to autoclaving. 4. Hazardous biochemicals, such as potential carcinogens, are to be carefully placed into a sealed container and stored in a fume hood prior to their disposal according to the institutional policy.

CLEANING AND PREPARATION OF CLEANING SOLUTION THE NEED FOR CLEANING: i. To remove the stains in the glassware. ii. To remove the chemical residues. iii. To remove the microbes partially by using the cleaning solution. iv. To remove the impurities which were stick on the surfaces of the glass wares. v. To remove the greasy areas by using mild detergents.

7

PREPARATION OF CLEANING SOLUTION AIM: To prepare the cleaning solution to clean the glass wares. REQUIREMENTS: Balance, erylyn Meyer flask (1 L), measuring cylinder, spatula, butter paper, potassium dichromate, conc sulphuric acid, metal distilled water etc. COMPOSITION FOR CLEANING SOLUTION: DILUTE SOLUTION CONC.SOLUTION Potassium dichromate 60 g 60 g. Water 1 litre 300 ml. Conc.sulphuric acid 60 ml 460 ml. PROCEDURE: About 800 ml of distilled water contained in a clean erylyn Meyer flask was dissolved with 60 gms of potassium dichromate and mixed with 60 gms of potassium dichromate and mixed with 200 ml of conc. H2SO4 and made upto a volume of 1 litre with distilled water. This solution is allowed to cool and used later. This cleaning solution which can be used to oxidise any organic matter and will clean the glassware like test tubes, petriplates, pipettes etc. The cleaning solution can be used until it terms into dark green colour solution. CLEANING: The glassware are soaked in the cleaning solution overnight and washed with soap water and rewashed in running tap water and then with distilled water. Glassware were allowed to drain of water and dried in a hot air oven at 800C for 2 hrs for further use.

8

Experiment No: STERILIZATION TECHNIQUES

Sterilization is the process of destroying all forms of microbial life-vegetative and sporulating. It is important that all equipment used in a microbiological experiment are sterilized in order that a particular organism of interest is grown, without contamination by the organisms present in the surrounding environment. Several methods of this sterilization are employed in the process of sterilization to sterilize the various equipments used in an experiment these methods are broadly classified as PHYSICAL and CHEMICAL METHODS OF STERILIZATION. I PHYSICAL AGENTS: The major physical agents used for the control of microorganisms are TEMPERATURE, RADIATION AND FILTRATION. 1. TEMPERATURE: Microorganisms can grow over a range of temperatures, from very low temperatures characteristic of Psychrophiles, to very high temperatures characteristic of thermophiles. Temperatures above a maximum generally kill microbes. Such high temperatures can be produced by DRYHEAT or MOIST HEAT. DRY HEAT STERILIZATION: Can be used where it is either undesirable or unlikely that steam under pressure will make direct and complete contact. This is true of glassware such as Petriplates, erlyn meyer flasks, pipettes, test tubes. Such material can be sterilized by placing in hot air, at a temperature of 1600C for 120 min. Alternatively, equipment like forceps, inoculation needle etc can be sterilized by direct heating on a flame till red hot. Thus it brings about destruction of unwanted organisms without changing the nature (flavour) of the material. This process involves heating at that temperature for 15 min and then cooking it quickly to 0- 50C. PRINCIPLE INVOLVED IN DRY HEAT STERILIZATION: Dry heat oxidise chemical components of organisms thus destroying them. MOIST HEAT STERILIZATION: High temperatures combined with high moisture is one of the most effective ways of sterilization. PRINCIPLES INVOLVED IN MOIST HEAT STERILIZATION: Moist heat coagulates microbial proteins, and is hence more rapid in killing microbes. Moist heat can be applied in the following ways in order to bring about sterility. STEM UNDER PRESSURE: Provides temperatures higher than those obtainable by any other method. 2t has advantages of rapid heating, penetration and moisture which facilitates coagulation of proteins. Autoclave is, a device used in the laboratory to sterilize media solution and to kill discarded cultures. It is operated at 15lbs/ sq inch pressure, which yields a temperature of 1210C effective in bringing about sterility in 15 min FRACTIONAL STERILIZATION OR TYNDALLISATION: Some microbial solutions cannot be heated over 1000C without being damaged. Such materials are sterilized by Tyndallization, which involves heating at 1000C on three successive days with incubation periods in between. Resistant spores germinate during the period of incubation that is killed on heating on the subsequent day. PASTEURISATION: Milk, cream and other alcohol beverages are subjected to controlled heat treatment which kills microbes of a certain type alone. 2. RADIATION: When ionising radiation pass through cells, they create free hydrogen radicals, hydroxyl radicals and peroxides that cause intracellular damage, resulting in destruction of microbes. This method of sterilization is effective for sterilization heat labile material / also called (OLD STERILIZATION)

9

U.V.light is the most effective region of the electromagnetic spectrum, and is employed in disinfecting of inoculation chamber and hospital operating rooms. UV light alters nucleic acids and results in a pyrimidine diamer, thus inhibiting DNA replication. 3. FILTRATION: This technique is used when the material to be used is heat labile and cannot be sterilized by heating for eg. Solutions of proteins, vitamins etc. filters of pore size 0.02μ -0.08μ are used to filters off microbes, thus rendering the filtrate sterile. Pore size, electric charge of the filter, charge carried by the organisms and nature of fluid being filtered affect efficiency of filtration e.g. Acity filters, Berkefeld filter, Berkefeld filter, membrane filter are all microbial filters. II CHEMICAL AGENTS Several groups of chemicals can be used as antimicrobial agents: S.No GROUP ACTION EXAMPLE 1. 2. 3. 4. 5. 6. 7.

Alcohol Aldehyde Halogens Heavy metals Gases Detergents Phenols

Denature proteins and solubilize lipids Alkylate, reacts with – NH2, SH- Co oH I2 inactivates proteins oxidise cells. Precipitates and inactivates proteins (used for surface sterilization) Alkylates organic compounds Disrupt cell membrane Denature protein and disrupt cell membrane

Ethyl Glutaraldehyde I2,Cl2 Hg Cl2 Ethylene dioxide

Ethyl alcohol (70%) finds indispensable use for disinfecting hands before and after a microbiological experiments, and also to disinfect the inoculation chamber or area where the experiment is conducted. CONCLUSION: Sterilization is the first indispensable step of any microbiological experiment. Clean and sterile equipment are pre-requisites for culture isolation and characterisation of any microorganism in a laboratory. Several methods can hence be employed to sterilize the various materials required for an experiment.

10

Experiment No:

PREPARATION OF MEDIA PURPOSE: To prepare the nutrient agar medium for bacterial culture. COMPOSITION OF NUTRIENT AGAR MEDIUM: [pH (7.2)] Beef extract 20g 1L Peptone 20g 1L Nacl 10 g 1L Agar 20g 1L Distilled water 1L COMPOSITION OF ROSE BENGAL AGAR MEDIUM pH: 5.6 ± 0.2 at 25 °C. (g/litre) Peptone 5.0; glucose 10.0; potassium dihydrogen phosphate 1.0; dichloran 0.002; magnesium sulfate 0.5; Rose Bengal 0.025; agar-agar 15.0. Distilled water 1L PROCEDURE: A clean erlynmeyer flask was taken with 200 ml of distilled water. The chemicals were weighed accurately and dissolved one by one taking care to add the next chemical only after the dissolution of the first one, expect agar and streptomycin in the case of rose bengal medium agar was melted separately and added to other ingredients. The pH was adjusted to 7.2 for nutrient agar medium and 6.5 for rose bengal agar medium. The total volume of the medium was made upto 1 litre. The medium was then distributed into 250 ml. Erlynmeyer flasks of 100 ml each and were plugged tightly with paper (from) and tied with thread. Then the flasks were sterilized at 1210C for 15 minutes with 15lbs/in2 in an autoclave. Streptomycin was added only after the sterilization of the rose bengal agar medium. PREPARATION OF PLATES, SLANTS AND DEEPS: The prepared media can be poured into plates, slants and deeps for cultures and pure-cultures. PLATES: The sterilized medium is cooled to 45 –500C and poured into sterile petriplates. About 20 ml of the medium is poured into each of the sterile petriplates and allowed to solidify. Incubate in an incubator overnight before use. SLANTS: The molten medium after adjusting the pH is poured into text tubes, 5 ml of each and the test tubes are placed in a slanted position so that they with solidify with maximum surface area. DEEPS: Deeps are prepared by pouring the molten medium in sterile glass text tubes and then placed in a vertical position. The tubes are then cooled in cold water and are used to maintain cultures for long periods of incubation. RESULT: Thus nutrient agar and rose bengal agar media were prepared and used to prepared and used to prepare plates, slants and deeps for inoculation.

Experiment No: ISOLATION, ENUMERATION AND PURIFICATION OF MICROBES FROM A GIVEN SAMPLE PURPOSE: To isolate, enumerate and purify various microbes from the given sample. PRINCIPLE The techniques commonly used for isolation of discrete colonies initially require that the number of organisms in the inoculum be reduced. The resulting diminution of the population size ensures that, following inoculation, individual cells will be sufficiently far apart on the surface of the agar medium to effect a separation of the different species present. The serial dilution is used to accomplish this. There are three techniques to do isolation of pure cultures. REQUIREMENTS: Sterile blanks (9ml), 12 sterile test tubes for slants, 1 sterile blank (10 ml), samples, 18-20 Petriplates(20 ml), 10 sterile pipettes (1 ml), bacterial growth medium- NUTRIENT AGAR (5 X 100 Ml), inoculation loop and wire, burner, marker pen, sterile chamber. PROCEDURE: 1. SERIAL DILUTION: Exactly 1 ml of the given sample was added to 9ml of blank,. This solution was labeled as 10-1. From this test tube, 1 ml of solution was taken and added to a 9 ml blank, mixed evenly and labeled as dilution 10-2 for the same manner, the procedure was separated during sterile pipettes for each transfer, until dilutions upto 10-5

are obtained. For the given soil sample, dilutions upto 10-3 and 10-4 were taken for fungal isolation and dilutions 10-4 were taken for fungal isolation and dilutions 10-4 and 10-5 were fungal isolation and dilutions 10-4 and 10-5 were taken for bacterial isolation.

2. ISOLATION OF MICROBES : POUR PLATE METHOD. Pour plating is a technique useful for isolation and enumeration of microbes 1 ml of the selected dilutions were pipetted into sterile petriplates in close proximity to the flame. Molten agar (nutrient agar in case of bacterial culture and Rose bengal agar in case of

11

12

fungal culture) was poured over the inoculum and the plates were swirled to evenly distribute the inoculum. Plates of a particular dilution were prepared in duplicates. The agar was allowed to solidify and the plates were incubated at a temperature of 350C in an inverted fraction for a period of 24-72 hours. SPREAD PLATE METHOD The spread-plate technique requires that a previously diluted mixture of microorganisms be used. During inoculation, the cells are spread over the surface of a solid agar medium with a sterile, L-shaped bent rod. The step-by-step procedure for this technique is as follows: Place the bent glass rod into the beaker and add a sufficient amount of 95% ethyl alcohol to cover the lower, bent portion.

a. With a sterile loop, place a loopful of culture in the center of the appropriately labeled nutrient agar plate that has been placed on the turntable. Replace the cover.

b. Remove the glass rod from the beaker and pass it through the Bunsen burner flame, with the bent portion of the rod pointing downward to prevent the burning alcohol from running down your arm. Allow the alcohol to burn off the rod completely. Cool the rod for 10 to 15 seconds.

c. Remove the Petri dish cover and spin the turntable. d. While the turntable is spinning, lightly touch the sterile bent rod to the surface

of the agar and move it back and forth. This will spread the culture over the agar surface.

e. When the turntable comes to a stop, replace the cover. Immerse the rod in alcohol and reflame.

f. Keep the plate for incubation 3. ENUMERATION OF MICROBES OBSERVATIONS AND CALCULATIONS BACTERIAL CIULTURE

The number of bacteria in a given sample is usually too great to be counted directly. However, if the sample is serially diluted and then plated out on an agar surface the number of colonies can be used as a measure of the number of viable (living) cells in that known dilution. However, keep in mind that if the organism normally forms multiple cell arrangements, such as chains, the colony-forming unit may consist of a chain of bacteria rather than a single bacterium. In addition, some of the bacteria may be clumped together. Therefore, when doing the plate count technique, we generally say we are determining the number of Colony-Forming Units (CFUs) in that known dilution. By extrapolation, this number can in turn be used to calculate the number of CFUs in the original sample.

Normally, the bacterial sample is diluted by factors of 10 and plated on agar. After incubation, the number of colonies on a dilution plate showing between 30 and 300 colonies is determined. A plate having 30-300 colonies is chosen because this range is considered statistically significant. If there are less than 30 colonies on the plate, small errors in dilution technique or the presence of a few contaminants will have a drastic effect on the final count. Likewise, if there are more than 300 colonies on the plate, there will be poor isolation and colonies will have grown together.

Generally, one wants to determine the number of CFUs per milliliter (ml) of sample. To find this, the number of colonies (on a plate having 30-300 colonies) is multiplied by the number of times the original ml of bacteria was diluted (the dilution factor of the plate

13

counted). For example, if a plate containing a 1/1,000,000 dilution of the original ml of sample shows 150 colonies, then 150 represents 1/1,000,000 the number of CFUs present in the original ml. Therefore the number of CFUs per ml in the original sample is found by multiplying 150 x 1,000,000 as shown in the formula below:

DILUTION P1 P2 AVG 10-4 10-5

No of bacterial C.F.U No of colonies Dilutions factor x amount of sample added = C.F.U / ml of sample. 4. STREAK PLATING: The streak-plate method is a rapid qualitative isolation method. It is essentially a dilution technique that involves spreading a loopful culture over the surface of an agar plate. Although many types of procedures are performed, the four-way, or quadrant, streak is described. Refer to figure, which schematically illustrates this procedure. Streaking cultures for isolated colonies allows you to:

• separate mixed cultures • purify a single type of bacterium • propagate a clonal population of bacteria • help with the identification of a bacterium

Streak plating is of two types: Quadrant streak:

a. Place a loopful of culture on the agar surface in Area 1. Flame and cool the loop and drag it rapidly several times across the surface of Area 1.

b. Reflame and cool the loop and turn the petri dish 900. Then touch the loop to a corner of the culture in Area 1 and drag it several times across the agar in Area 2. The loop should never enter Area 1 again.

c. Reflame and cool the loop and again turn the dish 900.Streak Area 3 in the same manner as Area 2.

d. Without reflaming the loop, again turn the dish 900 and then drag the culture from a corner of Area 3 across Area 4, using a wider streak. Don't let the loop touch any of the previously streaked areas. The flaming of the loop at the points indicated is to effect dilution of the culture so that fewer organisms are streaked in each area, resulting in the final desired separation.

SLANTS AND STABS: For long term storage of bacterial culture, suspended in a slow state of growth, slants and states can be used. Desired bacterial colonies were picked up with the sterile loop and streaked onto the surface of the slant (maximising the surface area) sterile inoculation were dipped into bacterial culture maintained in nutrient broth, and were stabbed into agar both slants and states were incubated. OBSERVATIONS: Purified and stored bacterial cultures were deserved and various types of colony morphology studied. SIZE: SHAPE/MARGIN: ELEVATION: PIGMENTATION: TEXTURE: RESULT: The given soil sample contained -------- C.F.U of bacteria per ml of sample. Desired bacterial cultures more purified, their morphology studied and subcultured for storage.

14

Experiment No: USE OF MICROSCOPE

A microscope is a scientific instrument with one or more lenses that allow you to observe specimens so small it is not visible to the naked eye, e.g. microorganisms (bacteria) and microsopic materials placed on the stage

General Procedures

1. Make sure all backpacks and junk are out of the aisles. 2. Plug your microscope in to the extension cords. Each row of desks uses the same cord. 3. Always start and end with the Scanning Objective. Do not remove slides with the high power objective into place - this will scratch the lens! 4. Always wrap electric cords and cover microscopes before returning them to the cabinet. Microscopes should be stored with the Scanning Objective clicked into place. 5. Always carry microscopes by the arm and set them flat on your desk.

Focusing Specimens

1. Always start with the scanning objective. Odds are, you will be able to see something on this setting. Use the Coarse Knob to focus, image may be small at this magnification, 15

16

but you won't be able to find it on the higher powers without this first step. Do not use stage clips, try moving the slide around until you find something.

2. Once you've focused on Scanning, switch to Low Power. Use the Coase Knob to refocus. Again, if you haven't focused on this level, you will not be able to move to the next level.

3. Now switch to High Power. (If you have a thick slide, or a slide without a cover, do NOT use the high power objective). At this point, ONLY use the Fine Adjustment Knob to focus specimens.

4. If the specimen is too light or too dark, try adjusting the diaphragm.

5. If you see a line in your viewing field, try twisting the eyepiece, the line should move. That's because its a pointer, and is useful for pointing out things to your lab partner or teacher.

Drawing Specimens

1. Use pencil - you can erase and shade areas 2. All drawings should include clear and proper labels (and be large enough to view details). Drawings should be labeled with the specimen name and magnification. 3. Labels should be written on the outside of the circle. The circle indicates the viewing field as seen through the eyepiece, specimens should be drawn to scale - ie..if your specimen takes up the whole viewing field, make sure your drawing reflects that.

Cleanup

1. Store microscopes with the scanning objective in place. 2. Wrap cords and cover microscopes. 3. Wash slides in the sinks and dry them, placing them back in the slide boxes to be used later. 4. Throw coverslips away.

17

Experiment No: SIMPLE STAINING

PURPOSE To perform the simple staining procedure to compare morphological shapes and arrangements of bacterial cells. PRINCIPLE In simple staining, the bacterial smear is stained with a single reagent. Basic stains with a positively charged chromogen are preferred, because bacterial nucleic acids and certain cell wall components carry a negative charge that strongly attracts and binds to the cationic chromogen. The purpose of simple staining is to elucidate the morphology and arrangement of bacterial cells. The most commonly used basic stains are methylene blue, crystal violet, and carbol fuchsin. MATERIALS Cultures 24-hour nutrient agar slant cultures of Escherichia coli and Bacillus cereus, and a 24-hour nutrient broth culture of Staphylococcus aureus. Reagents Methylene blue, crystal violet, and carbol fuchsin. Equipment Bunsen burner, inoculating loop, staining tray, microscope, lens paper, bibulous paper, and glass slides. PROCEDURE 1. Prepare separate bacterial smears of the organisms following the procedure described. Note: All smears must be heat fixed prior to staining. 2. Place a slide on the staining tray and flood the smear with one of the indicated stains, using the appropriate exposure time for each: Carbol fuchsin, 15 to 30 seconds; methylene blue, 1 to 2 minutes. 3. Wash the smear with tap water to remove excess stain. During this step, hold the slide parallel to the stream of water; in this way you can reduce the loss of organisms from the preparation. 4. Using bibulous paper, blot dry but do not wipe the slide. 5. Repeat this procedure with the remaining two organisms, using a different stain for each. 6. Examine all stained slides under oil immersion. OBSERVATIONS AND RESULTS In the space provided

1. Draw a representative field for each organism 2. Describe the morphology of the organisms with reference to their

shapes(bacilli,cocci,spirilli) and arrangements(chains,cluster,pairs).

18

Experiment No:

GRAM STAINING PURPOSE: To become familiar with 1. The chemical and theoretical bases for differential staining procedures. 2. The chemical basis of the Gram stain. 3. Performance of the procedure for differentiating between the two principle groups of bacteria: gram-positive and gram-negative. PRINCIPLE Differential staining requires the use of at least three chemical reagents that are applied sequentially to a heat-fixed smear. The first reagent is called the primary stain. Its function is to impart its color to all cells. In order to establish a color contrast, the second reagent used is the decolorizing agent. Based on the chemical composition of cellular components, the decolorizing agent may or may not remove the primary stain from the entire cell or only from certain cell structures. The final reagent, the counterstain, has a contrasting color to that of the primary stain. Following decolorization, if the primary stain is not washed out, the counterstain cannot be absorbed and the cell or its components will retain the color of the primary stain. If the primary stain is removed, the decolorized cellular components will accept and assume the contrasting color of the counterstain. In this way, cell types or their structures can be distinguished from each other on the basis of the stain that is retained. The most important differential stain used in bacteriology is the Gram stain, named after Dr.Christian Gram. It divides bacterial cells into two major groups, gram-positive and gram-negative, which makes it an essential tool for classification and differentiation of microorganisms. The Gram stain uses four different reagents. Descriptions of these reagents and their mechanisms of action follow. Primary stain Crystal Violet This violet stain is used first and stains all cells purple. Mordant Gram's iodine This reagent serves as a mordant, a substance that forms an insoluble complex by binding to the primary stain. The resultant crystal violet-iodine (CV-I) complex serves to intensify the color of the stain, and all the cells will appear purple-black at this point. In gram-positive cells only, this CV-I complex binds to the cell wall. The resultant magnesium-ribonucleic acid-crystal violet-iodine (Mg-RNA-CV-I) complex is more difficult to remove than the smaller CV-I complex. Decolorizing Agent Ethyl Alcohol, 95% This reagent serves a dual function as a lipid solvent and as a protein-dehydrating agent. Its action is determined by the lipid concentration of the microbial cell walls. In gram-positive cells, the low lipid concentration is important to retention of the Mg-RNA-CV-I complex. Therefore, the small amount of lipid content is readily dissolved by the action of the alcohol, causing formation of minute cell wall pores. These are then closed by alcohol's dehydrating effect. As a consequence, the tightly bound primary stain is difficult to remove, and the cells remain purple. In gram-negative cells,the high lipid concentration found in outer layers of the cell wall is dissolved by the alcohol, creating large pores in the cell wall that do not close appreciably on dehydration of cell wall

19

proteins. This facilitates release of the unbound CV-I complex, leaving these cells colorless or unstained. Counterstain Safranin This is the final reagent, used to stain red those cells that have been previously decolorized. Since only gram-negative cells undergo decolorization, they may now absorb the counterstain. Gram-positive cells retain the purple color of the primary stain The preparation of adequately stained smears requires that you bear in mind the following precautions: 1. The most critical phase of the procedure is the decolorization step, which is based on the ease with which the CV-I complex is released from the cell. Remember that over-decolorization will result in loss of the primary stain, causing gram-positive organisms to appear gram-negative. Under- decolorization, however, will not completely remove the CV-I complex, causing gram-negative organisms to appear gram-positive. Strict adherence to all instructions will help remedy part of the difficulty, but individual experience and practice are the keys to correct decolorization. 2. It is imperative that slides be thoroughly washed under running tap water between applications of the reagents. This removes excess reagent and prepares the slide for application of the subsequent reagent. 3. The best Gram stained preparations are made with fresh cultures , that is, not older than 24 hours. As cultures age, especially in the case of gram-positive cells, the organisms tend to lose their ability to retain the primary stain and may appear to be gram-variable; that is, some cells will appear purple, while others will appear red. Materials Cultures 24-hour nutrient agar slant cultures of Escherichia coli, Staphylococcus aureus, and Bacillus cereus. Reagents Crystal violet, Gram's iodine, 95% ethyl alcohol, and safranin. Equipment Bunsen burner, inoculating loop or needle, staining tray, glass slides, bibulous paper, lens paper, and microscope. PROCEDURE The steps are pictured in Figure 1. Obtain four clean glass slides. 2. Using sterile technique, prepare a smear of each of the three organisms and on the remaining slide prepare a smear consisting of a mixture of S.aureus and E.coli. Do this by placing a drop of water on the slide and then transferring each organism separately to the drop of water on the slide with a sterile, cooled loop. Mix and spread both organisms by means of a circular motion of the inoculating loop. 3. Allow smears to air dry and then heat fix in the usual manner. 4. Flood smears with crystal violet and let stand for 1 minute. 5. Wash with tap water. 6. Flood smears with the Gram's iodine mordant and let stand for 1 minute. 7. Wash with tap water.

20

8. Decolorize with 95% ethyl alcohol. Caution: Do not over-decolorize. Add reagent drop by drop until crystal violet fails to wash from smear. 9. Wash with tap water. 10. Counterstain with safranin for 45 seconds. 11. Wash with tap water. 12. Blot dry with bibulous paper and examine under oil immersion. OBSERVATIONS AND RESULTS Following your observation of all slides under oil immersion, record your results in the chart. 1. Make a drawing of a representative microscopic field. 2. Describe the cells according to their morphology and arrangement. 3. Describe the color of the stained cells. 4. Classify the organism as to the gram reaction: Gram-positive or gram-negative. Refer to photo numbers 2-4 in the color-plate insert for illustration of this staining procedure.

21

Experiment No:

SPORE STAINING (SCHAEFFER-FULTON METHOD)

PURPOSE:

To become familiar with 1. The chemical basis of the spore stain. 2. Performance of the procedure for differentiation between bacterial spore and vegetative cell forms.

PRINCIPLE Members of the anaerobic genera Clostridium and Desulfotomaculum and the aerobic genus Bacillus are examples of organisms that have the capacity to exist either as metabolically active Vegetative cells or as highly resistant, metabolically inactive cell types called Spores. When environmental conditions become unfavorable for continuing vegetative cellular activities, particularly with the exhaustion of a nutritional carbon source, these cells have the capacity to undergo sporogenesis and give rise to a new intracellular structure called the endospore, which is surrounded by impervious layers called spore coats. As conditions continue to worsen, the endospore is released from the degenerating vegetative cell and becomes an independent cell called a spore. Because of the chemical composition of spore layers, the spore is resistant to the deleterious effects of excessive heat, freezing, radiation, desiccation, and chemical agents, as well as to the commonly employed microbiological stains. With the return of favorable environmental conditions, the free spore may revert to a metabolically active and less resistant vegetative cell through germination. It should be emphasized that sporogenesis and germination are not means of reproduction but merely mechanisms that ensure cell survival under all environmental conditions. In practice, the spore stain uses two different reagents.

Primary Stain Malachite Green Unlike most vegetative cell types that stain by common procedures, the spore, because of its impervious coats, will not accept the primary stain easily. To further penetration, the application of heat is required. After the primary stain is applied and the smear is heated, both the vegetative cell and spore will appear green.

Decolorizing Agent Tap Water Once the spore accepts the malachite green, it cannot be decolorized by tap water, which removes only the excess primary stain. The spore will remain green. On the other hand, the stain does not demonstrate a strong affinity for vegetative cell components; the water removes it, and these cells will be colorless.

Counterstain Safranin This contrasting red stain is used as the second reagent to color the decolorized vegetative cells, which will absorb the counterstain and appear red. The spores retain the green of the primary stain.

MATERIALS Cultures

22

48- to 72-hour nutrient agar slant culture of Bacillus cereus and thioglycollate culture of Clostridium butyricum.

Reagents Malachite green and safranin. Equipment Bunsen burner, hot plate, staining tray, inoculating loop, glass slides, bibulous paper, lens paper, and microscope.

PROCEDURE 1. Obtain three clean glass slides. 2. Make individual smears in the usual manner using sterile technique. 3. Allow smear to air dry, and heat fix in the usual manner. 4. Flood smears with malachite green and place on a warm hot plate, allowing the preparation to steam for 2 to 3 minutes. Caution: Do not allow stain to evaporate; replenish stain as needed. Prevent the stain from boiling by adjusting the hot plate temperature. 5. Remove slides from hot plate, cool, and wash under running tap water. 6. Counterstain with safranin for 30 seconds. 7. Wash with tap water. 8. Blot dry with bibulous paper and examine under oil immersion.

OBSERVATIONS AND RESULTS Following your observation of all slides under oil immersion, record your results in the chart. 1. Make drawings of a representative microscopic field of each preparation. 2. Describe the location of the endospore within the vegetative cell as being central, subterminal, or terminal on each preparation. 3. Indicate color of the spore and vegetative cell on each preparation. Refer to photo number 6 in the color-plate insert for illustration of this staining procedure.

23

Experiment No: HANGING DROP TECHNIQUE FOR DEMONSTRATING MOTILITY OF BACTERIA PURPOSE: To determine bacterial motility

PRINCIPLE: Live, unstained, motile cells will demonstrate there motility under 400x magnification by displaying individual movement amongst cells. Meaning that movement is unique and directional. The hanging drop is used to distinguish this from brownian movement which is just a random jiggling motion. REQUIREMENTS: 12 hour-old broth culture of Proteus vulgaris Hanging drop (cavity) slide Cover slips Vaseline / Pertoleum jelly Match stick Wax marking pencil. PROCEDURE: 1. Clean and flame a hanging-drop slide and place it on the table with the depression uppermost. 2. Spread a little Vaseline or petroleum jelly around the cavity of the slide. 3. Clean a cover slip apply petroleum jelly on each of the four corners of the cover slip, using a match stick. 4. Place the cover lip on a clean paper with the petroleum jelly slide up. 5. Transfer one loopful of culture in the center of the cover slip. 6. Place the depression slide on to the coverslip, with the cavity facing down so that the depression covers the suspension. 7. Press the slide gently to form a seal between the cover slip and the slide. 8. Lift the preparation and quickly turn the hanging drop preparation cover slip up so that the culture drop is suspended. 9. Examine the preparation under low-power objective with reduced light. 10. Switch to the high-power objective and examine the preparations again. 11. Place a drop of oil on the cover slip and examine the preparation under oil-immersion objective. Precautions 1. If a bacterial growing on a solid medium is to be examined, a loopful of culture should be mixed with a drop of 2% CMC, in the center of the cover slip. 2. The depression slide is inverted over the cover slip insuch a way athat the suspension does not touch the surface of the concavity at any point. 3. The slide and cover slip should be sterilized after the examination is finished. 4. Petroleum jelly from the depression slide and cover slip should be removed at the end of the experiment, with xylene. OBSERVATION Make drawings of a representative microscopic field of each preparation

24

Experiment No: PREPARATAION OF BUFFERS AND MEASUREMENT OF pH PURPOSE: To prepare the buffers of required pH PRINCIPLE: The pH meter measures the electrical potential developed by a pair of electrodes dipping into a solution. For the measurement of pH of an electrode system sensitive to changes in the hydrogen ion activity of the solution is chosen. This electrode system consists of a sequence of electrodes whose potential varies with the pHof the solution. 1. Acetic acid-sodium acetate buffer: Reagents: 1. Acetic acid (0.2m) 1.55 ml of glacial acetic acid is made up to 100 ml with distilled water. 2. Sodium acetate solution (0.2m) 1.62 g of sodium acetate 0r 2.72 g of sodium acetate trihydrate is dissolved in 100 ml of water. PROCEDURE: Pipetted out exactly 35.2ml of sodium acetate solution into a 100ml standard flask. To this added exactly 14.8ml of acetic acid. This is made up to 100ml with distilled water. This gives 0.2m acetic acid-sodium acetate buffer whose pH could be measured a pH meter. The pH meter was first standardised using standard buffer. Washed the electrode with distilled water. Now introduced 02.m acetate buffer the pH was found to be.5. 2. Carbonate-Bicarbonate Buffer. Reagents:

i) Sodium Carbonate solution (0.2m) Dissolved 2.12 g of anhydrous sodium carbonate in 100 ml of water.

ii.Sodium Bicarbonate Solution (0.2M) Dissolved 1.68 g of sodium bicarbonate in 100 ml of water. PROCEDURE: Pipetted out exactly 27.5ml of sodium carbonate solution (0.2M). To this added 22.5ml of sodium bicarbonate solution (0.2M). This gives 0.2M carbonate-bicarbonate buffer, Whose pH is measured. Standardized the pH measured the pH of the prepared buffer. The pH was found to be 10. 3. Barbitone Buffer: REAGENTS: 1.Diethyl barbituric acid 2.Sodium diethyl barbiturate

PROCEDURE: Dissolved 2.589g of diethyl barbituric acid and 14.2 g of sodium diethyl barbiturate in distilled water and made up 1 liter. This gives borbitone buffer, Its pH was found to be 8.6 4. Citric acid Buffer: Reagents: Citric acid solution (0.1M) Dissoloved 2.101 g citric acid in 100 ml of water. PROCEDURE: Mixed 46.5ml of citric acid (0.1M) and 3.5 ml of sodium citrate (0.1M) and made upto 100 ml with water. This gives 0.1M citric acid buffer. Standardized the pH meter and measure the prepared solution, this give citrate buffer of pH 3. 5. Phosphate Buffer: Reagents:

1. Monobasic sodium phosphate solution (0.2M) Dissloved 2.78 g at monobasic sodium phosphate in 100ml of water

2. Dibasic sodium phosphate solution (0.2M) Dissloved 5.365g of dibasic sodium phosphate heptahydrate and 7.179 g of dibasic sodium phasphate deco carbonate in 100 ml of water. PROCEDURE: Mixed 39ml of monobasic sodium phosphate solution with 61ml of dibasic sodium phosphaste solution. This is made up to 200 ml with distilled water. This gives phosphate buffer whose pH can be measured. The pH of this was found top be 7. DIGITAL pH METER

RESULT: The buffers of the required pH was prepared 25

26

Experiment No: ESTIMATION OF REDUCING SUGAR BY BENEDICT’S METHOD AIM: To estimate the amount of glucose present in the given unknown solution using benedict’s quantitative reagent. PRINCIPLE: Benedict’s quantitative reagent is a modification of qualitative. It contains copper sulphate, sodium acetate and sodium corbonate. It also contains potassium thio cyanate and small amount of potassium ferricyanide. The inclusion of acetate prevents the precipitation of copper carbonate by chelating Cu3+ion. The thiocyanate causes the precipitation of white cuprous thio cyanate rather than red cupric oxide. On reduction of Cu3+ ions which enables the end point of the titration ie., the transition from blue to white to be readily observable. Methylene blue will be used as an additional indicator. The small amount of potassium ferricyanide prevents the re-oxidation of copper. A non-stoicheometric reaction is on which does not follow a defined pathway and cannot be described by an equation either quantitatively or qualitatively. The reduction of Cu3+ ions by sugar is a non-stoicheometric equation and is only constant over a small range of sugar concentration. To obtain accurate results the volume of sugar added must be with in 6- 12 ml for 10 ml of benedict’s reagent. If the preliminary titre value Falls outside this range the sugar solution must be titrations are repeated. REAGENTS REQUIRED: 1. STANDARD GLUCOSE SOLUTION: 200 mg of glucose was weighed accurately and made upto 100 ml with distilled water (concentration: 2 mg / ml) 2. BENEDICT’S QUANTITATIVE REAGENT 100 ml of solution acetate, 37.5 g of sodium carbonate and 62.5 g of potassium thiocyanate were dissolved in 300 ml of distilled water by warming gently and filtered. 9 g of copper sulphate is dissolved in 50 ml of water, added with continuous stirring. 2.5 ml of potassium ferricyanide is added and the volume is made upto 500 ml with water. 3. ANHYDROUS SODIUM CARBONATE PROCEDURE:

100 ml of benedict’s reagent was pipetted out into a clean conical flask. About 600 mg of anhydrous sodium carbonate was added to provide the required allcaling with a few porcelain bits and heated to boiling over a moderate flame. The standard glucose solution is taken in the burette when the benedict’s solution boils, glucose solution is added drop by drop (one drop per second) till the last trace of blue colour disappears. The volume of glucose rundown is noted and the titrations are repeated for concordant value. The given unknown sugar solution was made upto 100 ml in a standard flask with distilled water. Then the burette was filled with unknown sugar solution and the benedict’s reagent was titrated as before. The volume of sugar solution rundown was noted and titrations are repeated for concordant values. ESTIMATION OF REDUCING SUGAR BY BENEDICT’S METHOD

TITRATION 1 STANDARDISATION OF BENEDICT’S REAGENT BENEDICT’S REAGENT VS STANDARD GLUCOSE SOLUTION

S.No Volume of Benedict’s reagents(ml)

Burette Readings Volume of standard glucose(ml)

Indicator Intial ml

Final ml

Self

TITRATION 2: ESTIMATION OF GLUCOSE STANDARDISED BENEDICT’S REAGENT VS UNKNOWN GLUCOSE

S.No Volume of Benedict’s reagents(ml)

Burette Readings Volume of unknown glucose(ml)

Indicator Intial ml

Final ml

Self

CALCULATION: The standard glucose solution 2 mg / ml 5 ml of Benedict’s solution react with ml of the standard glucose solution. ml of standard glucose solution which contains x 2 = mg 5 ml of Benedict’s solution reacts with mg of unknown glucose 100 ml of unknown glucose contains is 100 x RESULT: The amount of glucose present in 100 ml of given unknown solution is Experiment No:

27ESTIMATION OF PROTEIN BY LOWREY’S METHOD

28

PURPOSE: To estimate the amount of protein present in the given solution PRINCIPLE: Protein in the given solution when treated with alkaline copper sulphate and Folin’s phenol reagent produces a blue colored complex. The intensity of the colour is directly proportional to the concentration of protein present in the given sample solution.

REAGENTS REQUIRED 1 Stock Solution: Bovine serum albumin of 100mg is weighed accurately and dissolved in 100ml of distilled water in a standard flask. (Concentration:1mg/ml) 2. Working Standard:

The stock solution of 10ml is diluted to 100ml with distilled water in a standard flask. (Concentration:100mg.ml) 3. Folin;s Phenol Reagent:

Folin’s phenol reagent is mixed with distilled water in a the ratio 1:2 4. Alkaline CuS04 Reagent: Solution A:

Sodium carbonate of 2% in 0.1N sodium hydroxide. Solution B: Sodium Potassium tartrate of 1% Solution C: Copper sulphate of 0.5% Solutions A,B,C are mixed in the proportion of 50:1:0.5 Unknown Preparation: The given protein is made upto 100ml with distilled water. PROCEDURE: Working standard of 0.2 to 1.0ml is pipetted out into clean test tubes labelled as S1, to S5 . Test solution of 0.2 and 0.4 ml is taken in test tubes labelled as T1 and T2. The volume is made upto 1.0ml with distilled water. Distilled water of 1.0ml serves as blank. To all the test tubes 4.5 ml of alkaline copper sulphate reagent is added and it is incubated at room temperature for 10 minutes. To all the test tubes 0.5ml of Folin’s Phenol reagent is added. The contents are mixed well and the blue colour developed is read at 640nm after 15 minutes. From the standard graph the amount of protein in the given unknown solution is calculated. ESTIMATION OF PROTEIN BY LOWREY’S METHOD

29

S.No Reagents B S1 S2 S3 S4 S5 T1 T2

1 Volume of working

standard (ml)

- 0.2 04. 0.6 0.8 1.0 - -

2 Concentration of working

standard (mg)

- 20 40 60 80 100 - -

3 Volume of unknown

solution (ml)

- - - - - - 0.2 0.4

4 Volume of distilled water

(ml)

1.0 0.8 0.6 0.4 0.2 - 0.8 0.6

5 Volume of alkaline

copper reagent (ml)

4.5 4.5 4.5 4.5 4.5 4.5 4.5 4.5

6 Volume of Folin’s phenol

reagent

0.5 0.5 0.5 0.5 0.5 0.5 0.5 0.5

The contents are mixed well and kept at room temperature for 10 minutes. The blue colour developed is

read at 640nm

7 Optical density 640nm

CALCULATION: of unknown solution corresponds to x OD

OD corresponds of y mg of protein

i.e. 0.2ml of unknown solution contains of protein

100ml of unknown solution contains

100 x ----

-----------

30

RESULT: The amount of protein present in the given solution is

Experiment No:

ESTIMATION OF CHOLESTEROL BY ZAK’S METHOD PURPOSE: To estimate the amount of cholesterol in an unknown food sample.

31

PRINCIPLE: Cholesterol in glacial acetic acid gives a red colour with ferric chloride and apolar sulphuric acid. This

reaction has been employed by ZAK’S to estimate the cholesterol in an unknown food sample.

REAGENTS REQUIRED: 1. Stock Standard Solution : About 100 mg of cholesterol was dissolved and made up to 100 ml with glacial acetic acid (concentration 1 mg / ml). 2. Working Standard : About 4 ml of stock solution was made up to 100 ml with ferric chloride acetic acid reagent (concentration in 40 mg / ml). 3. Ferric chloride of 0.05% in acetic acid. 4. Apolar sulphuric acid. 5. Glacial acetic acid. 6. Preparation of unknown food sample: 20 ml of food sample and 40 ml of chloroform was added and centrifuged. The supernatant was used for estimation. PROCEDURE: 0.5 ml to 2.5 ml of working standard were Pipetted out into a clean test tubes. About of 0.5 ml and 1 ml of unknown food sample supernatant was taken in a test tubes. The volume was made upto 5.0 ml with ferric chloride and 3.0 ml of concentrated sulphuric acid were added. The test tubes were kept at room temperature for 15 minutes. The pinkish red colour formed was measured at 540 nm. Standard graph was drawn for the values obtained. From the standard graph the amount of cholesterol present in the food sample can be calculated.

ESTIMATION OF CHOLESTEROL BY ZAK’S METHOD S.No

Reagents B S1 S2 S3 S4 S5 T1 T2

1. Volume of standard chloresterol (ml)

_ 0.5 1.0 1.5 2.0 2.5 _ _

2. Concentration of cholesterol (mg)

_ 20 40 60 80 100 _

_

3. Volume of food sample supernatant (ml)

_ _ _ _ _ _ 1

1

4. Volume of 0.05% ferric chloride acetic acid reagent (ml)

5 4.5 4 3.5 3 2.5 _ _

5. Volume of cone sulpheric acid (ml)

3

3 3 3 3 3 3

3

Incubated the tubes for 15 minutes at room temperature

6. O.D at 540 nm CALCULATION: 0.5 ml of standard corresponds to O.D 0.03 O.D corresponds to mg. ∴0.5 ml of unknown corresponds 100 ml of unknown corresponds to

32

RESULT: The amount of cholesterol present in unknown food sample was found to be Experiment No: ESTIMATION OF DNA BY DIPHENYLAMINE METHOD PURPOSE: To estimate the amount of DNA present in the given unknown solution. PRINCIPLE:

The deoxyribose of DNA in the presence of acid forms hydroxyl levolinic aldehyde which reacts with diphenylamine to give a blue colour. But only the deoxy ribose of purine nucleotide react. Reagents required: 1. Stock standard solution: Weighed 100 mg of DNA and dissolved in 100 ml of distilled water. 2. Working standard solution: 10 ml of stock was diluted to 100 ml using distilled water. 3. Diphenyl amine reagent: It should be prepared freshly by dissolving 1 gm of diphenyl amine in 100 ml of glacial acetic acid and by adding 2.5 ml of concentrated sulphuric acid. PROCEDURE: Pipetted out 0.2 ml – 1.0 ml of DNA solution into a series of test tubes and made up the volume to 3.0 ml with distilled water. 0.2 ml and 0.4 ml of unknown is taken and made upto 3.0 ml with water. Added 5.0 ml of disphenylamine reagent, mixed well and is heated in a boiling water bath for 10 minutes. Cooled and the colour developed is read at 595 nm. ESTIMATION OF DNA BY DIPHENYLAMINE METHOD

S.No Contents B S1 S2 S3 S4 S5 U1 U2

1. Volume of working standard in (m1)

- 0.2 0.4 0.6 0.8 1.0 - -

2. Concentration in μg - 20 40 60 80 100 - -

3. Volume of unknown in (m1) - - - - - - 0.2 0.4

4. Volume of distilled water in (ml)

3 2.8 2.6 2.4 2.2 2.0 2.8 2.6

5. Volume of diphenyl amine reagent in (ml)

5 5 5 5 5 5 5 5

Heated in a boiling water bath for 10 minutes

6. Optical density at 595 mm Calculation: 0.2 ml of unknown corresponds to 0.02 O.D ---O.D corresponds to ---- 0.2 ml of unknown corresponds to ---- 100 ml of unknown corresponds to = 100 X RESULTS: The amount of DNA present in 100 ml of given unknown solution

33

34

Experiment No: SEPARATION OF AMINO ACIDS BY THIN LAYER CHROMATOGRAPHY PURPOSE: To separate amino acids by thin layer chromatography. PRINCIPLE: Chromatography is a method by which a mixture of substances in smaller quantities can be separated both qualitatively and quantitatively. In chromatography there are two

35

phases-the stationary phase and other mobile phase. When the mobile phase moves along stationary phase, separation of substances takes place. In thin layer chromatography, the thin layer of gel functions as an inert supporting material. When the mobile phase moves along the gel solvent, as the partition coefficient differ for different sugars, the rate of flow differs and therefore separation occurs. Materials Required: 1.Silica gel G 2.Microscopic slides 3.n-butanol 4.Acetic acid

5.Spraying reagent (0.3% solution of Ninhydrin in butanol containing 3 ml acetic acid)

6.Amino acids PROCEDURE:

A slurry of silicagel G is prepared in 0.02M sodium acetate buffer. Taken the microscopic slides and keeping them flat, pipetted out about 1-2 ml of the slurry into them. By tilting the slides spread the slurry evenly on the surface. Lining the edges with Vaseline will be of help. Allowed the slides to dry completely leaving them flat. 5μl samples of amino acid (or mixture) are spotted and the slide is then dipped in a trough containing n-butanol-acetic acid-water in the ration 8:2:2. The slide must be handled with care not to break the surface. After development, that is, when the solvent has reached the top, the slide is dried and sprayed with the developing reagent. The slide is then heated in an oven at 1100 C for 10 minutes and Rf values of the spots are measured.

Rf = Distance moved by solute ---------------------------------

Distance moved by solvent RESULT:

Experiment No: SEPARATION OF SUGARS BY PAPER CHROMATOGRAPHY PURPOSE: To separate the sugars by paper chromatography. PRINCIPLE: Chromatography is a method by which a mixture of substances in smaller quantities can be separated both qualitatively and quantitatively. In chromatography there are two pahses the stationary phase and other mobile phase. When the mobile phase moves along

stationary phase, separation of substances takes place. In paper chromatography the paper functions as an inert supporting material. When the mobile phase moves along the paper solvent, As the partition coefficient differ for different sugars the rate of flow differs and therfore separation occurs. MATERIALS REQUIRED:

1.Whatmann No :1 filter paper 2.N-butanol 3.Glacial acetic acid 4.Sugars 5.Spraying reagent-Alkaline permanganate spraying agent 0.1% KmnO4 in 2% sodium carbonate

PROCEDURE: 3 strips of whatman No 1 filter paper of size 12×2 cm are cut. A line is drawn at a distance of 2.5 cm from the base and a pencil mark made at the mid point. Sugar solutions of glucose and fructose at a concentration of 100 mg/10ml is prepared. The chromatography paper is placed on a box having a slit with lighting arrangment underneath the slit. Spotting is done on the paper using a micropipette. Care is taken to see that the spot does not spread beyond a particular diameter. 10ml each glucose and fructose are spotted on a paper A and B. To the strips a mixture is applied. The three strips are placed in three different boiling tubes each containing 5ml of n-butanol acetic acid, water in the ration 4:2:1. The boiling tubes are plugged with cotton, The paper are kept in a a pre-saturated chromatographic chamber and the solvent is allowed to rise. When the solvent front has reached three fours of the paper the strips are removed and air dried. It is then sprayed with the spraying agent and dried in hot air oven at 1000C. The sugars appeared as yellow spots in a purple background. The distances travelled by the solvent are mesured. The distance from the base line to the centre of each spot are measured, Rf values is then calculated as follows.

dbysolventcetravelleDisysugartravelledbDisRF

tantan

=

RESULT: The Rf values of sugar is found to be

36