F Y BSc. Bot. (Sem I) Skill Enhancement Courses (SEC)

Skill Enhancement Courses (SEC)

Name of the Course: Tools and Techniques in Plant Science (Practical)

60 Hours                                                          Credits: 02

Course Objectives (CO):

CO1: Learn to use basic laboratory instruments and staining techniques in plant science.

CO2: Develop skills in microscopy, microtechniques, and slide preparation.

CO3: Master separation techniques and biochemical analysis of plant metabolites.

CO4: Acquire skills in molecular techniques, soil analysis, and computer applications for data presentation.

Course Outcomes:

COO1: Demonstrate proficiency in handling laboratory instruments and performing staining techniques essential for plant studies.

COO2: Prepare and analyze plant slides using various microscopy techniques.

COO3: Perform separation and qualitative analysis of plant metabolites using chromatography and other biochemical tests.

COO4: Apply molecular and soil analysis techniques, and use computer tools for data visualization and presentation in plant science.

(Test and Assessment Question Bank for CO Attainment and Analysis: Students should attempt a minimum of 2 questions from each Course Objective (CO)).

Course Objective (CO)

Relevant Questions

CO1: Learn to use basic laboratory instruments and staining techniques in plant science.

1)      How would you correctly operate a colorimeter to measure the absorbance of a plant extract?

2)      Describe the purpose and use of a laminar air chamber in plant science experiments.

3)      What steps would you follow to stain a plant tissue sample using safranin?

4)      What is the difference between the use of an autoclave and an oven in laboratory settings?

5)      Explain how to set up a microscope and adjust the focus to observe a whole mount slide of a plant sample.

CO2: Develop skills in microscopy, microtechniques, and slide preparation.

1)      How do you prepare a squash slide of a plant root tip for microscopic observation?

2)      Describe the procedure for preparing a transverse section (T.S.) of a leaf and explain its importance.

3)      What are the steps to create a smear slide of pollen grains, and why is it used in plant studies?

4)      Which stains are commonly used in whole mounts, and what cellular structures do they highlight?

5)      How would you differentiate between a longitudinal section (L.S.) and a transverse section (T.S.) of a plant stem?

CO3: Master separation techniques and biochemical analysis of plant metabolites.

1)      Explain the process of separating amino acids using paper chromatography.

2)       How would you perform a qualitative test to detect carbohydrates in a plant sample?

3)      What is the principle behind thin-layer chromatography, and how can it be used to separate plant sugars.

4)       Describe how to test for the presence of alkaloids in a plant extract and interpret the results.

5)       How do you verify Beer and Lambert’s Law using a colorimeter and a plant pigment solution?

CO4: Acquire skills in molecular techniques, soil analysis, and computer applications for data presentation.

1)      Outline the steps involved in extracting genomic DNA from a plant leaf.

2)      How would you measure the pH of a soil sample, and why is this parameter important for plant growth?

3)      How would you use MS Excel to create a graph that displays the results of a soil pH test?

4)      Describe the procedure for determining the temperature of a soil sample and discuss its relevance to plant science.

5)      Explain how you would use photo micrographic techniques to document your findings from a microscopic examination.

List of Practical’s

Module 1: Introduction to Laboratory Tools and Instruments

1.1 Study of Basic Laboratory Instruments (Microscope, Colorimeter, Autoclave, Oven, Incubator, Laminar Air Chamber, Tilak Air Sampler).

1.2 Study of stains and staining techniques.

Module 2: Microscopy and staining Techniques

2.1 Microslide Preparation—Whole Mounts, Smears, Squashes

2.2 Plant Microtechnique (T.S., L.S., R.L.S., T.L.S.)

Module 3: Separation techniques

3.1 Separation of Amino Acids by Paper Chromatography

3.2 Separation of Sugars by Thin-Layer Chromatography

Module 4: Biochemical analysis

4.1 Qualitative Tests for plant Metabolites (Proteins, Carbohydrates, Alkaloids, Tannins)

4.2 Study and verification of Beer and Lambert’s Law

Module 5: Basic Molecular Technique

5.1 Genomic DNA Extraction

Module 6. Soil Analysis Techniques

6.1 Determination of temperature and pH of different Soil Samples

Module 7. Computer Techniques

71. Study of photo micrographic techniques

7.2. Use of computer for preparation of Tables, Graphs and Presentations (MS Office)

Note: This course is designed to provide foundational skills in plant science laboratory techniques. The practical assessments and final project are structured to evaluate students' proficiency and understanding, preparing them for potential careers in plant science research and related fields. 

Module 1: Introduction to Laboratory Tools and Instruments

1.1 Study of Basic Laboratory Instruments (Microscope, Colorimeter, Autoclave, Oven, Incubator, Laminar Air Chamber, Tilak Air Sampler).

 01: Microscope:

The microscope is a crucial instrument used to view small objects or details that are invisible to the naked eye by magnifying their images. It's particularly useful for observing the shapes of bacteria, fungi, parasites, and host cells in both stained and unstained samples.

a. Types of Microscopes

1. Simple Microscope: Utilizes a concave mirror and contains a single adjustment screw for focusing.

2. Compound Microscope: Uses a plane mirror on one side and a concave mirror on the other, featuring a coarse adjustment for rapid focusing. It's the most common type used in microbiology and consists of two lens systems for magnification. These microscopes can be monocular (single eyepiece) or binocular (two eyepieces).

b. Parts of the Microscope: 

The main parts of the microscope are the eye-pieces, microscope tube, nosepiece, objective, mechanical stage, condenser, coarse and fine focusing knobs, and light source.

 i.            Eye-pieces: The specimen is viewed through the eyepiece, which magnifies the image formed by the objective lens. The magnification power ranges from 5x to 20x. Binocular microscopes allow for the adjustment of the distance between eyepieces to fit the user's eyes.

 ii.            Microscope Tube: Attached to the top of the arm, this tube can be monocular or binocular and supports the eyepiece.

 iii.            Mechanical Tube Length: The distance between the objective lens insertion point and the top of the draw-tube for eyepieces.

 iv.            Nose-piece: Located under the arm, it holds and rotates the objectives, arranging them in order of increasing magnifying power to prevent contamination with immersion oil.

  v.            Objectives: These lenses, typically with magnifications of 4x, 10x, 40x, and 100x, form the initial image of the specimen. The numerical aperture (NA) indicates the lens’s light-gathering ability, correlating with its resolving power. Immersion oil, which has the same refractive index as glass, is used with the 100x objective to enhance image clarity. The 100x objective is for oil immersion.

The numerical aperture (NA) is the measure of light-gathering power of a lens. The NA corresponding to the various magnifying powers of the objective is:

Magnification                                           Numerical aperture

10x                                                              0.25

40x                                                             0.65

100x                                                        1.25

A high NA indicates a high resolving power and thus useful magnification

To provide the best image at high magnification, immersion oil is placed between the slide and the oil immersion objective (100x). Unlike air, immersion oil has the same refractive index as glass. Therefore, it improves the quality of the image. If immersion oil is not used, the image appears blurred or hazy

    vi.            Condenser: Illuminates the specimen and controls light intensity and contrast. Some condensers can be adjusted vertically using a rack-and-pinion mechanism.

  vii.            Two-sided Mirror: Reflects light to illuminate the specimen, with one plain side for artificial light and a concave side for natural light. It can rotate freely and is usually mounted at the base of the microscope.

c. Functioning of the Microscope

A compound light microscope relies on three primary optical components to produce a sharp and clear image: the condenser, the objectives, and the eyepieces. Each part plays a crucial role in the functionality of the microscope.

1. Condenser: The condenser is responsible for illuminating the specimen. It does this by converging a parallel beam of light onto the specimen from either a built-in or natural light source. This concentrated light enhances the visibility and contrast of the specimen, making it easier to observe fine details.

2. Objectives: The objective lenses are located close to the specimen and are the first to magnify the image. The objectives form a magnified, inverted (upside-down) image of the specimen. These lenses come in various magnifying powers, such as 4x, 10x, 40x, and 100x (oil immersion), allowing for different levels of detail to be observed.

3. Eyepieces: The eyepiece, or ocular lens, is where the viewer looks through to see the magnified image. It further magnifies the image created by the objective lens. The final image, which is observed by the viewer, is formed below the plane of the slide. Eyepieces typically have magnifications ranging from 5x to 20x.

The total magnification of the compound light microscope is determined by multiplying the magnifying powers of the objective lens and the eyepiece. For example, if the eyepiece has a magnification of 10x and the objective lens has a magnification of 100x, the total magnification is calculated as follows:

Total Magnification=Magnification of Eyepiece × Magnification of Objective

Total Magnification=10x×100x=1000x

This means the specimen will be magnified 1000 times its actual size, allowing for detailed observation of microscopic structures.

02 Colorimeter: A colorimeter is an instrumental device used in colorimetry to measure the absorbance of a specific wavelength of light by a sample solution. Invented by Louis J. Duboscq in 1870, this device is fundamental in determining the concentration of solutes in a solution by comparing the color intensity of the sample to that of a reference solution with a known concentration.

Instrumentation of Colorimeter

1.      Light Source:

1. • Provides energy across the visible spectrum (380-780 nm). Tungsten lamps are typical for visible and near-infrared ranges, while halogen deuterium lamps are used for the UV range (200-900 nm).

2.      Slit:

2. • Reduces stray light by allowing a precise beam to pass through.

3.      Condensing Lens:

3. • Produces a parallel beam of light after it passes through the slit.

4.      Monochromator:

o Filters monochromatic light from polychromatic light, allowing only the desired wavelength to pass through. Types include prism, glass, and gratings.

o Prism: Refracts light when it passes through.

o Glass: Selectively transmits specific wavelength ranges.

o Gratings: Made of graphite, separates light into different wavelengths.

5.      Cuvette (Sample Cell):

5. • Holds the sample solution. Sizes vary (square, rectangle, round) and typically have a 1 cm diameter. Types include glass, quartz, and plastic.

   • Glass Cuvettes: Absorb light at 340 nm, cost-effective.

   • Quartz Cuvettes: Allow both UV and visible light.

   • Plastic Cuvettes: Inexpensive, easily scratched, shorter lifespan.

6.      Photocell (Photodetector):

6. • Converts light energy into electrical energy, measuring light intensity.

7.      Galvanometer:

7. • Detects and measures the electrical signal from the photocell, displaying optical density (OD) and percentage transmission.

Principle of Colorimeter

When an incident light beam (I0) passes through a solution, part of it is reflected (Ir), absorbed (Ia), and the remainder is transmitted (It). The relationship is:

I0=Ir+Ia+It

By measuring the initial intensity (I0) and the transmitted intensity (It), the absorbance (Ia) can be determined, as the reflected light (Ir) is kept constant using cuvettes with identical characteristics.

Colorimeter Operating Procedure

        i.            Power On: Rotate the Power Switch knob clockwise. Allow the device to warm up for 15 minutes to stabilize the light source and detector.

      ii.            Set Wavelength: Adjust the Wavelength Control knob to select the desired wavelength for the measurement.

    iii.            Switch to Transmittance Mode: Press the MODE control key until the “Transmittance” indicator lights up.

    iv.            Zero Adjustment: Use the Zero Control knob to set the display’s T-factor to 0.0% with the sample chamber empty and the cover closed.

      v.            Insert Blank Solution: Fill a cuvette with the blank solution. Clean the exterior to remove any contaminants, then place it in the sample chamber, aligning and securing it properly.

    vi.            Calibrate Transmittance: Adjust the Transmittance/Absorbance control knob to set the display to 100.0% transmittance.

  vii.            Switch to Absorbance Mode: Press the MODE control key to switch to Absorbance mode. The display should read 0.0. Adjust if necessary.

viii.            Insert Sample Solution: Place the sample solution in another cuvette, clean the exterior, and insert it into the sample chamber.

    ix.            Read Measurements: Read the %T value directly from the display. Switch to Absorbance mode to read the A value.

      x.            Remove Cuvette: - Reverse the insertion process to remove the cuvette from the sample chamber.

Applications of Colorimeter

        i.            Printing Industry: Evaluates the quality of ink and paper by measuring color consistency and accuracy.

      ii.            Food Industry: Measures the composition of food products, including color additives and concentration of ingredients.

    iii.            Medical Labs: Analyzes biochemical samples such as blood, urine, and cerebrospinal fluid for various diagnostic purposes.

    iv.            Textile and Paint Industries: Assesses the quality and color of textile and paint products to ensure consistency and quality control.

03 Autoclave:

An autoclave is a device that uses steam under pressure to sterilize materials by killing bacteria, viruses, and spores. It is widely used in healthcare and various industries for its effective sterilization method, which leverages moist heat.

Autoclave Parts/Components

1. Pressure Chamber:

• The core component consisting of an inner chamber and an outer jacket.

• Inner chamber: Made of stainless steel or gunmetal.

• Outer jacket: Made of iron, filled with steam to accelerate the sterilization process.

• Sizes range from 100 to 3000 liters.

2. Lid/Door:

• Seals off the external atmosphere to create a sterile environment.

• Made airtight with scrsew clamps and an asbestos washer.

• Includes several key components:

  • Pressure Gauge: Indicates the pressure inside the autoclave.

  • Pressure Releasing Unit/Whistle: Controls pressure by releasing excess vapor.

  • Safety Valve: Releases pressure if it becomes dangerously high to prevent explosions.

3. Steam Generator/Electrical Heater:

• Located beneath the chamber, it heats water to generate steam.

• Water levels must be monitored to avoid damage to the heating system or interference with the contents.

4. Vacuum Generator (if applicable):

• Removes air from the chamber to prevent air pockets that could harbor microorganisms.

5. Wastewater Cooler:

• Cools the effluent before it enters the drainage system to prevent damage from boiling water.

Principle of Autoclave Operation

Autoclaves operate on the principle of moist heat sterilization. Steam under pressure increases the boiling point of water, allowing higher temperatures to be reached for effective sterilization. The steam penetrates materials, coagulating proteins and irreversibly inactivating microbes. The standard conditions are 121°C at 15 psi for 15 minutes, but this can vary based on the material's contamination level.

Autoclave Operating Procedure: Autoclaving typically involves running the machine at 121°C for at least 30 minutes using saturated steam at a minimum of 15 psi of pressure. Below is a step-by-step guide for running an autoclave:

1. Pre-Check: Ensure the autoclave is free of any items from the previous cycle.

2. Add Water: Fill the chamber with a sufficient amount of water.

3. Load Materials: Place the items to be sterilized inside the chamber.

4. Seal the Chamber: Close the lid securely and tighten the screws to create an airtight seal. Turn on the electric heater.

5. Adjust Safety Valves: Set the safety valves to maintain the required pressure.

6. Boil Water: Allow the water to boil. The air-water mixture should escape through the discharge tube to displace all the air inside. Complete displacement is indicated when water bubbles cease to emerge from the pipe.

7. Close Drainage Pipe: Once air displacement is complete, close the drainage pipe.

8. Reach Desired Pressure: Allow the steam to build up to the desired pressure level (typically 15 psi). The whistle will blow to release excess pressure once the desired level is reached.

9. Holding Period: Maintain the autoclave at the desired pressure and temperature for the holding period, usually 15 minutes.

10. Cool Down: Turn off the electric heater and let the autoclave cool until the pressure gauge shows that the pressure has dropped to atmospheric levels.

11. Release Pressure: Open the discharge pipe to allow air to enter and equalize the pressure inside the autoclave.

12. Unload Materials: Finally, open the lid and remove the sterilized items from the chamber.

Uses of Autoclave

1. Healthcare and Laboratories: Sterilizes medical equipment, glassware, surgical instruments, and medical waste.

2. Research and Development: Sterilizes culture media, autoclavable containers, plastic tubes, and pipette tips.

3. Waste Management: Inactivates regulated medical waste containing biological materials before disposal.

4. Industry: Used in pharmaceuticals, food processing, and other industries to ensure sterile conditions.

04 Hot Air Oven:

A hot air oven is essential laboratory equipment used for sterilizing objects and samples through dry heat. This method, known as dry heat sterilization, was introduced by French scientist Louis Pasteur in the late 1800s. Pasteur used dry heat briefly to eliminate harmful microorganisms in wine without altering its taste.

Hot air ovens are ideal for sterilizing heat-resistant items that do not melt, change form, or catch fire at high temperatures. They effectively kill microorganisms and bacterial spores at high temperatures over several hours. Effective sterilization requires selecting the appropriate temperature and holding time based on the type of microorganism and material. Common settings are 170°C for 30 minutes, 160°C for 60 minutes, and 150°C for 150 minutes.

Principle of Hot Air Oven

Hot air ovens operate on the principle of dry heat sterilization using convection, conduction, and radiation. Heating elements heat the air inside the chamber, which is evenly circulated with the help of fans. This hot, dry air exposure heats the external surfaces of items, transferring heat inward. In microorganisms, this heat evaporates water, causing oxidative damage to cellular components, protein denaturation, and ultimately cell death.

Components of a Hot Air Oven

Mechanical Parts:

1. Coat/Cabinet: Made of aluminum or stainless steel, providing insulation and preventing heat loss.

2. Fibreglass: Insulates the space between the outer cabinet and inner chamber, using brown or yellow fiberglass.

3. Chamber: Rectangular chamber made of aluminum or stainless steel with ribs for shelves.

4. Shelves (Mesh): Aluminum plates that hold items, allowing air movement and aeration.

5. Motorized Fans/Blower: Distributes hot air evenly inside the chamber.

6. Door: Single door with asbestos gasket to reduce heat loss.

Electrical Parts:

1. Power Supply: 220V-50Hz transformer and rectifier.

2. Heater: Generates heat through electrical current, with various heater types operating from 50 to 300°C.

3. Thermostat: Heat sensor directly connected to the heater, preventing temperature overshoot.

4. Temperature Indicator: Thermometer or thermocouple to measure internal temperature.

5. Timer: Electrical or mechanical timers for setting sterilization duration.

6. Fuse: Prevents electrical damage during short circuits or high loads.

7. Control Panel: Allows control of temperature, time, with indicator lamps and switch knobs.

Hot Air Oven Operating Procedure

1. Power On: Plug in and switch on the oven.

2. Preheat: Preheat the oven for 30 minutes before loading items.

3. Set Temperature and Time: Adjust the temperature gauge based on the content volume.

4. Load Items: Place items on shelves with adequate spacing for heat circulation.

5. Close Door: Fasten screws and let the temperature rise.

6. Monitor Temperature: Check the thermometer to ensure the desired temperature is reached.

7. Sterilization Period: Once the holding period is achieved, switch off the oven and allow it to cool.

8. Unload Items: Remove items using oven mitts or tongs, then close the door.

Applications of Hot Air Oven

1. Laboratory Sterilization: Sterilizes glassware, metal items, culture media, and non-volatile compounds.

2. Quality Testing: Tests food items, pharmaceuticals, and consumables for temperature stability.

3. Research Use: Used in biology, chemistry, and material science research.

4. Sample Treatment: Heat treatment and drying of metals, alloys, soil, and other materials.

05 Incubator:

An incubator in microbiology is an insulated, enclosed device that maintains optimal temperature, humidity, and other environmental conditions necessary for the growth of organisms. It is crucial for cultivating both unicellular and multicellular organisms under controlled conditions.

Components of an Incubator

Cabinet: The main body of the incubator, a double-walled cuboidal structure, with a capacity ranging from 20 to 800L. The outer wall is made of stainless steel, while the inner wall is aluminum, with glass wool insulation in between to prevent heat loss and reduce energy consumption. Inward projections on the inner wall support shelves inside the incubator.

Door: The insulated door includes a glass panel for observing the interior without opening it, and a handle for easy maneuvering.

Control Panel: Located on the outer wall, the control panel contains switches and indicators for controlling the incubator's parameters, including a switch for the thermostat.

Thermostat: Used to set and maintain the desired temperature inside the incubator.

Perforated Shelves: Shelves with perforations to allow hot air circulation, which can be removable for cleaning.

Asbestos Door Gasket: Provides an airtight seal between the door and cabinet to maintain the internal environment.

L-shaped Thermometer: Positioned on the outer wall with one end inside the chamber to read the internal temperature.

HEPA Filters: Advanced incubators may include HEPA filters to reduce contamination from airflow.

Humidity and Gas Control: CO2 incubators have a water reservoir for maintaining humidity and gas chambers for CO2 concentration control.

Principle and Working of an Incubator

An incubator operates on the principle that microorganisms require specific environmental conditions for growth. It maintains optimal conditions of temperature, humidity, oxygen, and CO2 levels.

• Temperature Control: The thermostat regulates temperature using heating and no-heating cycles. The thermometer allows external temperature monitoring.

• Insulation: Insulation maintains an isolated environment inside the cabinet, promoting effective microbial growth.

• Humidity and Airflow: Controlled through various mechanisms to simulate natural growth conditions.

• CO2 Concentration: Adjustable to balance pH and humidity for optimal organism growth.

• Shaking Incubators: Variations include shaking incubators for continuous culture movement, aiding cell aeration and solubility studies.

Operating Procedure

1.      Preparation: Ensure no leftover items from previous cycles are in the incubator. If cultivating multiple organisms with the same requirements, they can be placed together.

2.      Preheating: Close the door, switch on the incubator, and heat to the desired temperature. Use the thermometer to confirm the temperature.

3.      Parameter Setting: Set required CO2 concentration and humidity levels if needed.

4.      Loading Cultures: Place petri dish cultures upside down on perforated shelves to prevent condensation on the medium surface.

5.      Sealing: For long incubation periods, seal plates with adhesive tape or place them in plastic containers.

6.      Incubation: Lock the door and maintain the cultures inside for the required period.

Applications of Incubators

1.  Microbial and Cell Culture: Growing and maintaining microbial or cell cultures.

2.  Growth Rate Acceleration: Enhancing the growth rate of slow-growing organisms.

3.  Biochemical Oxygen Demand: Supporting microbial colony reproduction for biochemical oxygen demand tests.

4.  Breeding and Hatching: Used in zoology for breeding insects and hatching eggs.

5.  Controlled Storage: Providing a controlled environment for sample storage before laboratory processing.

06 Laminar Air Chamber (Laminar Air Flow):

A laminar flow hood, also known as a laminar air flow cabinet, is an enclosed workstation designed to create a contamination-free environment through the use of filters that capture all particles entering the cabinet. It is particularly useful for aseptic media distribution and plate pouring.

Components:

i. Cabinet:

• Constructed from stainless steel to prevent spore collection, with minimal gaps or joints.

• Insulated to maintain an uncontaminated environment inside.

• Features a glass shield on the front, which may either fully open or have openings for hand access.

ii. Working Station:

• A flat surface inside the cabinet where processes occur.

• Made of stainless steel to prevent rusting and facilitate easy cleaning.

• Used to place culture plates, burners, loops, and other necessary tools.

iii. Filter Pad/Pre-filter:

• Located at the top, it traps dust particles and some microbes before the air enters the cabinet.

iv. Fan/Blower:

• Positioned below the filter pad, it sucks in and circulates air within the cabinet.

• Directs air towards the HEPA filter to trap remaining microbes.

v. UV Lamp:

• Sterilizes the interior of the cabinet and its contents before use.

• Should be turned on 15 minutes before operation to avoid exposure to UV rays.

vi. Fluorescent Lamp:

• Provides adequate lighting inside the cabinet during operation.

vii. HEPA Filter:

• Ensures a sterile environment by trapping fungi, bacteria, and dust particles.

• Provides particulate-free air by filtering pre-cleaned air.

Principle/Working of Laminar Flow Hood

The laminar flow hood operates based on the principle of maintaining a laminar flow of air through the cabinet.

• Air is drawn in through one or more HEPA filters, creating a particulate-free environment.

• The filtration system processes the air, which then flows uniformly across the work surface.

• Air first passes through the pre-filter, ensuring streamlined flow into the cabinet.

• The blower directs air towards the HEPA filters, which trap contaminants, allowing only clean air to flow out.

• Effluent air exits through perforations at the bottom rear of the cabinet and over the workbench towards the operator’s face.

• The sides of the hood are enclosed, and positive air pressure is maintained to prevent external contaminated air from entering.

Procedure for Running the Laminar Flow Cabinet:

1. Preparation: Ensure no items sensitive to UV rays are inside the cabinet.

2. Sterilization: Close the glass shield and switch on the UV light for about 15 minutes to sterilize the workbench.

3. Airflow: Switch off the UV light and wait for about 10 minutes before turning on the airflow.

4. Operation: Start the airflow approximately 5 minutes before beginning work. Open the glass shield and switch on the fluorescent light. Optionally, sterilize the workbench with disinfectants like 70% alcohol for additional protection.

5. Completion: After completing the work, turn off the airflow and fluorescent lamp, and close the glass shield.

Uses of Laminar Flow Cabinet

1. Laboratory Applications: Used for contamination-sensitive processes like plant tissue culture. Ideal for media plate preparation and organism culture.

2. Pharmaceutical Industry: Used in drug preparation techniques to ensure a contamination-free environment.

3. Specialized and General Lab Techniques: Can be custom-made for specific tasks or used for general microbiological and industrial lab procedures.

07: Tilak’s Continuous Air Sampler:

Tilak’s Continuous Air Sampler is an innovative and indigenous device designed by Prof. S.T. Tilak and Kulkami in 1970. This air sampler earned Prof. Tilak the President’s Medal awarded by the Invention Promotion Board, New Delhi, in 1972.

Equipment Description and Working

Technical Specifications:

• Dimensions: 10.4” x 10.4” x 8”

• Power Supply: AC 230V

• Sampling Rate: 5 Litres/minute (0.17 Ft³/minute)

• Sampling Duration: Continuous for 7 days

Components:

• Cubical Tin Box: The main body of the sampler.

• Elevated Round Cap: Placed over the closing lid at the top.

• Electric Clock: Synchronizes with the rotating drum.

• Projecting Tube: Sucks air through an orifice at 5 Litres/minute.

• Rotating Drum: Coated with a thin layer of petroleum jelly on cellotape, it captures bio components from the air. The drum completes one full rotation in 7 days.

• Fan: A small fan with three prongs fixed in the cover creates negative pressure by forcing air out of the collection chamber.

• Exhaust Area: Measures 6 x 2.7 cm, located in the lid.

Working Principle:

• Air is drawn into the sampler through the projecting tube, impinging on the cellotape coated with petroleum jelly on the rotating drum.

• This setup entraps biocomponents as the drum rotates, providing a continuous trace of air samples over a week.

• The collected samples are then mounted on slides using glycerine jelly for microscopic examination.

Applications:

• Agricultural Research: Used for monitoring airborne spores in crop fields, aiding in disease management and optimizing fungicide applications.

• Environmental Monitoring: Assesses air quality and bioaerosols in various ecosystems.

• Public Health: Studies allergenic spores and pollen, contributing to allergy and respiratory disease management.

• Industrial Use: Ensures clean air environments in pharmaceutical and manufacturing settings.

• Academic Research: Supports studies in aerobiology and aeromycology.

Module 1: Introduction to Laboratory Tools and Instruments

1.2 Study of stains and staining techniques.

 Aim: To study Stains and Staining techniques.

Stains: A stain is a substance used in microscopy and various biological applications to enhance the visibility of cells, tissues, and their components. 

Staining: Staining is a crucial technique in microscopy used to enhance contrast in microscopic images. It involves using stains and dyes to view biological tissues more clearly, often with the aid of different microscopes.

Purposes of Staining

·         Enhance Visualization: 

·         Highlight Metabolic Processes:

·        Examine Bulk Tissues and Cell Populations: 

· 

Fixation: Fixation involves several steps aimed at preserving the shape of cells or tissues as accurately in its original stage as much as possible. Heat fixation is sometimes used to kill the cells, adhere them to a slide, and make them permeable to stains.

Mordant: A mordant is a chemical that enhances the binding of a dye to a substance, such as cells or tissues, by forming an insoluble compound with the dye.

Accentuators: Accentuators are substances that enhance the selective staining power, leading to more intense and vivid staining.

Mounting: Mounting is the process of preparing samples for microscopic observation by attaching them to a glass microscope slide.

Staining Techniques

• Direct (Positive) Staining: In this technique, the organism itself is stained while the background remains unstained.

• Indirect (Negative) Staining: Here, the background is stained, leaving the organism unaltered and visible against the stained background.

Kinds/Types of Staining Technique:

Stains are categorized into three main types:

1.      Simple Stain

2.      Differential Stain

3.      Acid-Fast Staining  (Ziehl–Neelsen stain)

1. Simple Staining:

Simple staining involves immersing a sample in a dye solution, followed by rinsing and observation. Some dyes require a mordant, a chemical that reacts with the dye to form an insoluble, colored precipitate, which helps the stain remain fixed even after washing away excess dye. This method typically uses only one dye and can be performed with basic dyes in direct staining or acidic dyes in negative staining. Simple staining is useful for studying cellular morphology, examining the nature of cellular contents, and determining the intracellular location of cell.

Commonly Used Simple Stains:

·         Methylene Blue: Applied by flooding the smear with the stain for 2 minutes, then allowing it to air dry. This stain clarifies the morphology of organisms such as Fungi,  H. influenzae and Gonococci.. Stains acidic cell parts (like nucleus) blue. Used on animal, bacteria and blood specimens. Can be used as a substitute for Janis B green.

·         Dilute Carbol Fuchsin: Prepared by diluting carbol fuchsin with distilled water to a ratio of 1/15. The smear is stained for 30 seconds, then washed and blotted dry. This stain is used for throat swabs, as a counterstain in Gram staining, and to highlight the morphology of Vibrio cholerae.

·         Polychrome Methylene Blue: Produced by allowing Loeffler’s Methylene Blue to ripen slowly in half-filled bottles, which is aerated periodically. The stain develops a polychrome property through slow oxidation, enhanced by the addition of potassium carbonate. This stain is used to demonstrate the McFadyean reaction of B. anthracis, showing blue bacilli surrounded by purple capsular material.

2. Differential Stains:

Differential stains utilize multiple dyes to categorize cells into different groups or types based on their structural and chemical characteristics. Unlike simple stains, which mainly reveal cell morphology, differential staining provides more detailed information about cell wall properties, such as thickness.

Gram Staining is a key differential staining technique used to differentiate bacterial species into two main categories: Gram-positive and Gram-negative. This method, developed by Hans Christian Gram, involves using crystal violet as the primary dye, iodine as a mordant, and a counterstain like safranin or fuchsin.

Principles of Gram Staining:

·         Crystal Violet: Stains all bacterial cell walls.

·         Gram’s Iodine: Acts as a mordant to fix the primary dye.

·         Decolorizer: Removes crystal violet from Gram-negative bacteria, which have a thinner peptidoglycan layer and an outer membrane.

·    

Gram Reaction:

·         Gram-Positive Bacteria: Retain the crystal violet stain due to their thick peptidoglycan layer and appear dark blue or violet.

·         Gram-Negative Bacteria: Lose the crystal violet stain during decolorization and take up the counterstain, appearing red or pink. Their cell walls are thinner with an additional outer membrane.

Gram Stain Procedure

1. Prepare the Slide: Place a drop of NaCl solution onto a glass microscope slide.

2. Sample Collection: Using a sterilized and cooled inoculation loop, collect a small amount of bacterial colony.

3. Mixing: Gently mix the bacteria into the NaCl drop on the slide.

4. Air Drying: Allow the bacterial sample to air dry completely.

5. Heat Fixing: Using a slide holder, pass the dried slide through the flame of a Bunsen burner 3 to 4 times, with the smear side facing up.

6. Primary Stain: Flood the slide with crystal violet stain and let it sit for 1 minute.

7. Rinse: Rinse the slide gently with water to remove excess stain.

8. Mordant Application: Apply iodine to the slide, covering it completely, and let it sit for 1 minute.

9. Rinse: Rinse the slide with water to remove excess iodine.

10. Decolorization: Flood the slide with acetone alcohol. After 10 to 15 seconds, rinse with water. Avoid overexposure to prevent removing the crystal violet from Gram-positive cells.

11. Secondary Stain (Counterstain): Flood the slide with safranin and let it sit for 1 minute.

12. Rinse: Rinse the slide with water to remove excess safranin.

13. Drying: Allow the slide to air dry completely.

14. Microscopy: Place the stained smear on the microscope stage, smear side up. Apply a drop of immersion oil directly to the smear and examine using the 100X objective lens.

3. Acid-Fast Staining:

The Ziehl–Neelsen stain, commonly known as the acid-fast stain, is a widely used differential staining technique. This method was first described by two German doctors, bacteriologist Franz Ziehl (1859-1926) and pathologist Friedrich Neelsen (1854-1894). Some bacteria resist decolorization by both acid and alcohol, classifying them as acid-fast organisms. This technique divides bacteria into two groups: acid-fast and non-acid-fast, and is extensively used in diagnosing tuberculosis and leprosy. Mycobacterium tuberculosis, the causative agent of tuberculosis (TB), is the most significant .

Acid-Fast Staining (Ziehl-Neelsen) Procedure:

1. Prepare the Smear: Make a smear, allow it to air dry, and then heat fix it.

2. Primary Stain: Flood the smear with Carbol Fuchsin stain, a lipid-soluble, phenolic compound that can penetrate the cell wall.

3. Filter Paper: Cover the flooded smear with filter paper.

4. Steam: Steam the slide for 10 minutes, adding more Carbol Fuchsin as needed.

5. Cool and Rinse: Allow the slide to cool and rinse with deionized (DI) water.

6. Decolorize: Flood the slide with acid alcohol (3% HCl and 95% ethanol) or 20% H₂SO₄ for 15 seconds. Tilt the slide at a 45-degree angle over the sink and add acid alcohol dropwise until the red color stops streaming from the smear.

7. Rinse: Rinse the slide with DI water.

8. Counter Stain: Add Loeffler’s Methylene Blue stain to the smear, which will color non-acid-fast cells blue. Leave the stain on for 1 minute.

9. Final Rinse and Dry: Rinse the slide and blot it dry.

10. Microscopy: Use an oil immersion objective to view the stained slide.

Common Stains in Botany:

In plant science, specific stains are commonly used to highlight different tissues and structures. These stains should be handled with care, as they can be hazardous to health and can stain clothing and skin. Proper safety equipment such as gloves and lab coats should always be worn. Here are some commonly used stains in botanical experiments:

1. Safranin O (1%)

Formulation:

• 50 mg safranin powder

• 50 ml distilled water

Dissolve safranin powder in water while stirring, then filter. Safranin O stains xylem and fibers bright red, and nuclei and plastids pink. Mainly used for sections of plant tissues, stains red.

2. Light Green

Formulation:

• Light Green powder

• Distilled water or alcohol

Light Green stains plant tissues green, often used in conjunction with other stains to highlight different collagen fibers.

3. Fast Green

Formulation:

• Fast Green powder

• Distilled water or alcohol

Fast Green stains plant cell walls and connective tissues, often used in combination with other stains for detailed tissue analysis.

4. Acetocarmine

Formulation:

• Acetocarmine powder

• Acetic acid

Acetocarmine is used to stain chromosomes in dividing cells, highlighting them in a reddish color, making it useful for studying mitosis and meiosis.

5. Iodine Potassium Iodide (IKI, Lugol Solution)
Formulation:

o 2 g potassium iodide (KI)

o 100 ml distilled water

o 0.2 g iodine Dissolve potassium iodide in water, then add iodine. This solution stains starches blue to black.

Iodine Stains carbohydrates in plant and animal specimens brown or blue-black. Stains glycogen red.

Caution: Iodine vapors are toxic. Avoid inhalation and handle with care.

6. Safranin O (1%)
Formulation:

o 50 mg safranin powder

o 50 ml distilled water Dissolve safranin powder in water while stirring, then filter. Safranin O stains xylem and fibers bright red, and nuclei and plastids pink.

o Mainly used for sections of plant tissues, stains red.

7.1: Toluidine Blue O (TBO)
Formulation:

o 50 mg Toluidine Blue O

o 100 ml water Dissolve the powder in water and adjust the pH to 6.8. TBO is a metachromatic stain: lignin appears blue to blue-green, pectins pink, and nuclei blue to greenish-blue.

o Stains acidic cell parts (like nucleus) dark blue. Used to show mitosis in plant cells.

7.2: Aniline Blue
Formulation:

o 0.5% (w/v) aniline blue in 0.2 M phosphate buffer (pH 6.5) or deionized water Filter through Whatman no. 1 paper. Aniline blue stains callose tissue.

8. Phloroglucinol HCl
Formulation:

o 2 g Phloroglucinol

o 80 ml of 20% ethanol

o 20 ml concentrated HCl (12 M) Dissolve Phloroglucinol in ethanol, then add HCl. Cover with aluminum foil to prevent degradation. This stain highlights lignin.
Caution: Mix in a fume hood as concentrated HCl is very corrosive. Handle with care.

9.   Sudan VI, Sudan Black, Oil Red O, and Nile Blue:

These are Lipid staining involves using dyes soluble in lipids to visualize intracellular lipids in tissue sections. These dyes can be substituted for one another, as they all work on the principle that the dye is more soluble in the lipid than in the solvent.

Procedure:

1. Cut the sample into 8-10 micron sections and air dry.

2. Rinse with 60% isopropanol.

3. Stain with Oil Red O solution for 15 minutes.

4. Rinse with 60% isopropanol.

5. Dip in alum hematoxylin to stain the nuclei.

6. Rinse with distilled water.

7. Mount in water or glycerin jelly.

10 Coomassie Brilliant Blue (CBB)

Formulation:

• 0.1% Coomassie Brilliant Blue R-250 powder

• 50% methanol

• 10% acetic acid

• 40% distilled water

Preparation:

1. Dissolve 0.1 g of Coomassie Brilliant Blue R-250 in 50 ml of methanol.

2. Add 10 ml of acetic acid.

3. Add 40 ml of distilled water.

4. Mix well until the dye is completely dissolved.

Usage: Coomassie Brilliant Blue (CBB) stain is a widely used method for routine visualization of proteins separated on polyacrylamide gels. 

Module 2: Microscopy and staining Techniques

2.1 Micro slide Preparation—Whole Mounts, Smears, Squashes

(In Module 1, you learned about the working and handling of a compound microscope. Cells of all organisms are not visible to the naked eye and can only be observed under a microscope. In this exercise, you will learn some routinely used methods to prepare slides for microscopic observations.)

Aim: To Study Micro slide Preparation—Whole Mounts, Smears, Squashes

Requirement:

Step 1: Before preparing microscope slide, gather all the necessary instruments:

• Flat or Concave Slides

• Personal Protective Equipment (PPE) such as safety glasses and gloves

• Cover Slips (Optional)

• Distilled Water

• Sterile Forceps

• Sterile Pipette/Dropper

• Inoculation/Smear Loop

If you plan to perform a slide staining, you will also need:

• Stain Solution

• Staining Rack

• Blotting Paper

Methods of Slide Preparation:

1.      Whole Mounts

2.      Smears

3.      Squashes

1. Whole Mounts Method: This method is used for minute plant specimens (small size) that do not need cutting or sectioning, e.g., filamentous algae, algal thallus , small leaves, parts of the large leaf, stripped leaf and stem epidermis , hairs or pollen and parts of flower. To prepare a complete samples of higher plants, small leaves or small parts of large leaves or flower parts or stripped leaf and stems epidermises, and other parts. To study the characteristics of the epidermal cells, such as stomata, trichomes and cuticle.

Whole mount preparation is used for minute plant specimens that do not require cutting or sectioning, such as filamentous algae, small leaves, and parts of flowers. This method helps study the characteristics of epidermal cells, including stomata, trichomes, and cuticles. Whole mounts are categorized into three types: Temporary, Semi-permanent, and Permanent.

Temporary Whole Mount

• Purpose: Mainly for classwork.

• Method: Mount the plant sample in water.

Semi-permanent Whole Mount

• Duration: Materials are prepared for a few hours to a fortnight.

• Mounting Medium: Glycerin (pure or diluted) or glycerin jelly (Kaiser’s glycerin jelly, which consists of 1 part pure gelatin and 6 parts water).

• Advantage: Preserves the natural colors of plant specimens.

• Sealing: Slides are sealed with Canada balsam.

• Ideal for: Unicellular and colonial algae, mosses protonemata, fungal spores, and fern prothalli.

Permanent Whole Mount Preparation

Two commonly used methods are the Hygrobutol method and the Glycerine-Xylol method.

1.      Hygrobutol Method

1. • Fixation: Kill and fix the specimen in Chrome-acetic.

   • Washing: Wash the material in running water.

   • Staining: Use suitable stains like Harris haematoxylin, Iron Haematoxylin, or Mayer’s carmalum.

   • Differentiation: Wash the material in differentiating solutions (15%, 30%, 50%, and 70% ethyl alcohol) for at least 20 minutes in each.

   • Alcohol Treatment: Leave the specimen in 85% alcohol for 18 hours.

   • Counterstain: Dissolve in 95% alcohol and methyl cellosolve.

   • Combination: Primary stains (Haematoxylin) and counterstains (Erthyrosin B, orange G, Fast green).

   • Dehydration: Gradually add hygrobutol (90 parts hygrobutol and 10 parts ethyl alcohol).

   • Mounting: Transfer specimens to Butyl Balsam and allow evaporation for 2 hours, then mount permanently.

2.      Glycerine-Xylol Method

2. • Fixation: Kill and fix the specimen in Chrome acetic.

   • Washing: Thoroughly wash the material in running water.

   • Staining: Stain in Haematoxylin solution.

   • Glycerine Treatment: Transfer the specimen to 10% aqueous glycerine and leave undisturbed until it reaches the consistency of pure glycerine.

   • Dehydration: Use absolute alcohol for dehydration.

   • Xylol Treatment: Replace alcohol with xylol, then mount with balsam and cover slip for permanent preparation.

.

2.  Smear Method: The smear method aims to spread individual cells and cell organelles suspended in a liquid into a homogeneous single layer on a glass slide. This ensures that the cells or organelles are killed and fixed without artifacts. Since the cells or organelles stick to the slide, adhesives are unnecessary, avoiding pigmentation and artifacts. This method is used for preparing plant specimens with single-cell bodies, such as bacteria, some fungi, algae, isolated cells, or cell organelles like plastids and mitochondria. Smear slides are typically used for analyzing chromosomes, blood cells, seminal fluids, tissue culture, throat swabs, and differential staining of bacteria.

Procedure:

i.            Sample Application:

o Place a drop of the liquid sample (e.g., slime, or bacteria) on a slide.

ii.            Smearing:

o Using the edge of a second slide held at a 45° angle, slowly smear the sample to create a thin, even coating.

i.            Covering:

o Place a cover slip over the sample, ensuring no air bubbles are trapped.

ii.            Removing Excess Liquid:

o Remove any excess liquid with tissue paper.

iii.            Drying:

o Allow the smear to dry naturally in an environment of moderate, steady temperature. The angle of the smearing slide affects the length of the smear; a steeper angle results in a shorter smear. For blood samples, back the smearing slide into the sample and then push across the slide, pulling the blood in the opposite direction to create a smooth layer. Thicker slides can be created with two drops, but only with mammalian blood due to the lack of a nucleus in erythrocytes, allowing cells to amass in multiple layers.

iv.            Fixing (for bacterial samples):

o Fix the smear by passing the slide repeatedly over a flame.

v.            Observation:

o The smear can either be stained or observed directly.

o Place a cover slip over the sample, ensuring no air bubbles are trapped.

o Remove excess liquid with a tissue or the corners of a filter paper.

o Allow the slide to air dry for 2-3 minutes, then mount using DPX.

3.  Squash Method: The squash method is essential for studying the internal structures of botanical samples and their relationships, particularly in cases where sectioning does not suffice. It is used to observe spores within sporangia, meiosis in pollen mother cells, and mitosis in stem and root tips.

Applications:

• Examining mitosis and meiosis in plant tissues (e.g., onion root tips and buds)

• Studying cell division in animal tissues (e.g., grasshopper testes, Drosophila salivary glands)

Materials Required:

1. Plant Materials:

   • Onion root tips

   • Onion flower buds

2. Animal Tissues:

   • Testis of grasshopper

3. Chemicals:

   • Acetic alcohol (1 part acetic acid, 3 parts ethyl alcohol)

   • 2N hydrochloric acid (HCl)

   • Staining solutions (e.g., iodine, methylene blue, crystal violet, 1% aceto-carmine or acetoorcein)

   • 45% acetic acid

   • 70% alcohol

4. Equipment:

   • Slides

   • Cover slips

   • Pasteur pipettes

   • Spirit lamp

   • Watch glass

   • Filter paper

   • Dissection kit

   • Nail polish or DPX mountant

   • Compound microscope

Detailed Procedure for Plant Samples:

1.  Sample Preparation:

1. • Growing Onion Roots:

     • Place onion bulbs with the root side down in water in a beaker or conical flask.

     • Allow roots to grow for 3 days until they reach a length of 2-3 cm.

   • Fixation:

     • Cut the root tips (2-3 cm long).

     • Transfer root tips to a solution of acetic alcohol (1 part acetic acid, 3 parts ethyl alcohol) for 12-24 hours.

     • Store fixed root tips in 70% alcohol until they are used for squash preparation.

2.  Hydrolysis and Staining:

2. • Hydrolysis:

     • Transfer root tips from the fixative (acetic alcohol) to a watch glass and wash extensively with water.

     • Add a few drops of 2N HCl to the watch glass.

     • Hydrolyze the tissues in 2N HCl for 10 minutes at room temperature or for 1 minute over a spirit lamp flame. Ensure the watch glass is moved over the flame to prevent tissue damage.

   • Staining:

     • After hydrolysis, wash the root tips in water to remove HCl.

     • Add 1% aceto-carmine or acetoorcein stain to the root tips for 10-15 minutes.

3.  Squash Preparation:

3. • Preparation of Stained Root Tips:

     • Transfer 2-3 stained root tips onto a slide.

     • Cut and retain only the meristematic region, removing any debris.

   • Adding Acetic Acid:

     • Place a drop of 45% acetic acid on the meristematic region.

     • Carefully place a cover slip over the stained material.

     • Remove excess acetic acid from the edges of the cover slip using the edge of a filter paper.

   • Squashing the Sample:

     • Place the slide between two sheets of filter paper.

     • Gently press the cover slip down vertically with your thumb or the flat end of a pencil to flatten cells and spread chromosomes.

   • Sealing:

     • Seal the edges of the cover slip with nail polish to minimize fluid evaporation.

4.  Observation and Results:

4. • Microscopic Observation:

     • Place the slide on the microscope stage.

     • Observe the specimen under low magnification first, then under higher magnifications.

   • Identifying Mitosis Stages:

     • Prophase: Chromosomes appear as two chromatids with a single centromere; the nuclear membrane slowly disappears.

     • Metaphase: Chromosomes align at the cell's equatorial plate.

     • Anaphase: Chromosomes move to opposite poles of the cell.

     • Telophase: Chromosomes reach the poles, and the nuclear membrane re-forms.

Module 2: Microscopy and staining Techniques

2.2 Plant Micro technique (T.S., L.S., R.L.S., and T.L.S.)

 Aim : Study of different techniques of plant sectioning.

Objective: To prepare and observe transverse sections (T.S.), longitudinal sections (L.S.), radial longitudinal sections (R.L.S.), and tangential longitudinal sections (T.L.S.) of plant stems and roots using plant microtechnique.

Materials:

Plant stem (e.g., sunflower or maize)  

Microscope slides and cover slips

Microtome or sharp blade/razor

Fixative solution (e.g., FAA - Formalin-Acetic Acid-Alcohol)

Embedding material (e.g., paraffin wax/ Potato)

Staining solutions (e.g., Safranin and fast green)

Forceps and dissecting needles

Microscope (compound)

Procedure:

1. Sample Collection and Fixation:Collect fresh samples of a plant stem and root.Immediately place the samples in the fixative solution (FAA) and leave for 24 hours to preserve the tissue structure.

2. Dehydration and Embedding: After fixation, dehydrate the samples by passing them through an increasing concentration series of ethanol (70%, 85%, 95%, and 100%). Embed the dehydrated samples in melted paraffin wax/potato cavity and allow the wax to solidify (Fig. 1).

3. Sectioning:

a. Transverse Section (T.S.)

• Preparation:

o A transverse section (T.S.) is a cross-sectional slice taken perpendicular to the longitudinal axis of the plant organ (e.g., stem or root).

o Using a microtome or a sharp blade, thin sections of the plant stem or root are carefully cut across the diameter of the organ.

o These sections are then stained using a double-staining method, typically with safranin to highlight lignified tissues (e.g., xylem) and fast green to stain other tissues (e.g., phloem, cortex).

Application:

• Tissue Identification:

  • T.S. is crucial for observing the arrangement and distribution of various tissues such as epidermis, cortex, vascular bundles, and pith.

  • The method allows the identification of primary and secondary growth, differentiation between monocot and dicot plants, and observation of the vascular cambium in stems undergoing secondary growth.

• Understanding Vascular Structure:

  • In dicots, the T.S. reveals the characteristic arrangement of vascular bundles in a ring, with xylem towards the inside and phloem towards the outside.

  • In monocots, the vascular bundles appear scattered throughout the ground tissue.

• Comparative Anatomy:

  • It is used to compare different plant species or different organs (e.g., roots vs. stems) to understand their structural adaptations.

b. Longitudinal Section (L.S.)

Preparation:

• • A longitudinal section (L.S.) is a slice cut along the length of the plant organ, parallel to its longitudinal axis.

  • The section can be taken from the stem, root, or leaf. In the case of stems, the section should include the apical meristem to observe the elongation process.

  • After cutting, the sections are stained using the same staining method as T.S.

Application:

• Cell Elongation and Differentiation:

  • L.S. allows the observation of cell elongation, particularly in growing regions like the shoot or root apices. It shows how cells transition from the meristematic zone (actively dividing cells) to the zone of elongation and differentiation.

• Observation of Tissue Differentiation:

  • This section helps in understanding the development of various tissues from the apical meristem, including the formation of primary xylem, phloem, and ground tissue.

• Growth Patterns:

  • It is used to study growth patterns and the role of hormones in cell elongation and differentiation.

c. Radial Longitudinal Section (R.L.S.)

• Preparation:

  • A radial longitudinal section (R.L.S.) is a slice taken along the radius of the cylindrical organ (e.g., woody stem), extending from the center outward.

  • The section is made to pass through the center of the organ, allowing for observation of the tissues along a radial plane.

  • This section is stained similarly to T.S. and L.S.

Application:

• Secondary Growth Observation:

  • R.L.S. is essential for studying secondary growth in woody plants. It shows the radial arrangement of secondary xylem and phloem, providing insights into the growth rings and the age of the plant.

• Vascular Cambium Activity:

  • It allows for the examination of the activity of the vascular cambium and how it contributes to the formation of secondary xylem (wood) and secondary phloem.

• Wood Anatomy:

  • The section is used in wood anatomy to study the structure and arrangement of wood cells, including tracheids, vessels, and fibers, which are crucial for understanding wood properties and taxonomy.

d. Tangential Longitudinal Section (T.L.S.)

• Preparation:

  • A tangential longitudinal section (T.L.S.) is a slice taken parallel to the outer surface of the cylindrical organ (e.g., woody stem), tangential to the growth rings.

  • The section is made parallel to the tangential plane, often showing the outermost tissues.

  • The section is stained using the same methods as for other sections.

Application:

• Tissue Arrangement in Woody Plants:

  • T.L.S. is particularly useful for observing the arrangement of cells and tissues in the bark and outer wood layers. It reveals the tangential organization of vascular rays and phloem fibers.

• Growth Ring Analysis:

  • It is used to study growth rings and the orientation of wood fibers, helping in dendrochronology (the study of tree rings) and understanding environmental influences on growth.

• Cell Orientation:

  • The section provides insights into the orientation of cells within the wood, which is important for understanding mechanical properties and processing of wood in industries.

4. Staining: Place the sections in a staining solution (e.g., safranin) for a few minutes to stain the lignified tissues.Rinse the sections in water and counterstain with fast green to highlight other tissues. Dehydrate the stained sections using ethanol (95% and 100%) to remove excess stain and water.

5. Mounting: Place the stained sections on a microscope slide. Add a drop of mounting medium and carefully place a cover slip over the section to avoid air bubbles.

6. Microscopic Observation: Observe the prepared slides under the compound microscope at low and high magnifications. Identify and note the arrangement of different tissues (e.g., epidermis, cortex, vascular bundles) in each section. Sketch or take micrographs of the observed sections.

7. Recording Observations: For each type of section (T.S., L.S., R.L.S., T.L.S.), record the following:

• • T.S.: Arrangement and distribution of tissues such as xylem, phloem, and cortex.

  • L.S.: Elongation and differentiation of cells along the plant’s length.

  • R.L.S.: Radial arrangement of secondary xylem and phloem.

  • T.L.S.: Tangential organization of cells and tissues in woody stems.

8. Discussion: Compare the observations from different sections. Discuss the significance of each section in understanding plant structure and function. Reflect on any difficulties encountered during the sectioning or staining processes and suggest improvements.

Conclusion:

This experiment provides hands-on experience in preparing and analyzing various types of plant sections. It reinforces students' understanding of plant anatomy and the importance of microtechnique in studying plant tissues.

Module 3: Separation techniques

3.1 Separation of Amino Acids by Paper Chromatography

Aim: To Separation of Amino Acids by Paper Chromatography

Objective: To separate and identify amino acids present in a sample using paper chromatography and to calculate the Rf values for each amino acid.

Materials Requirements:

• Whatman No. 1 filter paper (10x20 cm)

• 600 mL beaker

• 2% aqueous ammonia in 20 mL 2-propanol (Solvent)

• 0.05 M aspartic acid, glycine, and tyrosine in 1.5% hydrochloric acid

• Capillary tubes

• Ninhydrin solution (2% in ethanol)

• Oven (100°C)

• Stapler

• Ruler

Procedure:

1. Preparation of the Filter Paper:

   • Draw a pencil line 2 cm from the bottom edge along the long axis of the filter paper.

   • Mark eight 'x' spots 2 cm apart on the line, starting 3 cm from the left edge.

2. Spotting the Amino Acids:

   • Use separate capillary tubes to place a small drop of each amino acid (aspartic acid, glycine, tyrosine) and your unknown sample on the marked spots.

   • Ensure each spot is no larger than 2 mm in diameter.

   • Allow the spots to air dry for 10 minutes.

3. Preparing the Chromatogram:

   • Roll the filter paper into a cylinder, with the pencil line on the outside.

   • Staple the ends of the paper to form a cylinder, ensuring the edges don’t touch each other.

4. Developing the Chromatogram:

   • Place the paper cylinder in the 600 mL beaker, ensuring that the baseline is above the solvent level but the paper does not touch the sides of the beaker.

   • Allow the solvent to move up the paper for 90 minutes or until it is 1 cm below the top edge.

5. Visualizing the Amino Acids:

   • After 90 minutes, remove the paper and mark the solvent front.

   • Allow the paper to dry.

   • Your instructor will spray the paper with ninhydrin solution.

   • Heat the chromatogram in an oven at 100°C for 10-15 minutes to develop the spots.

6. Calculating Rf Values:

   • Measure the distance each amino acid traveled from the baseline.

   • Measure the distance the solvent front traveled.

   • Calculate the Rf value for each amino acid using the formula: 

Conclusion:

By performing paper chromatography, successfully separate and identify amino acids in the sample and calculate their Rf values to compare with known standards..

 Rf1 value =………….. denoted that the sample are with amino acid………………..

 Rf2 value =………….. denoted that the sample are with amino acid………………..

Rf3 value =………….. denoted that the sample are with amino acid………………..

3.2 Separation of Sugars by Thin-Layer Chromatography

Aim:
To separate and identify sugars in a sample using Thin-Layer Chromatography (TLC)

Materials Required:

• TLC plates (silica gel)

• 0.5 mg bark extract (dissolved in solvent)

• Mobile phase solvent (e.g., butanol acid in a 12:3:5 ratio)

• Capillary tubes

Objective:

To separate sugars present in the bark extract using TLC and identify them through specific staining techniques.

Procedure:

1. Sample Preparation:

• Dissolve 0.5 mg of bark extract in an appropriate solvent (e.g., water or methanol).

• Prepare the TLC plate by drawing a baseline 1 cm from the bottom edge using a pencil.

• Use a capillary tube to apply small spots of the extract onto the TLC plate along the baseline. The spot should be small, around 1-2 mm in diameter.

2. Chromatographic Separation:

• Prepare the mobile phase solvent by mixing butanol, acetic acid, and water in a 12:3:5 ratio.

• Pour the prepared solvent into the chromatography chamber.

• Place the TLC plate into the chamber so that the solvent level is below the baseline where the sample was spotted.

• Allow the solvent to rise up the plate through capillary action until it is approximately 1 cm from the top edge of the TLC plate.

• Remove the plate from the chamber and allow it to air dry completely.

3. Visualization of Sugars:

• After the plate is completely dry, spray it lightly with 0.2% ninhydrin solution.

• Gently heat the plate (using a hot plate or oven) to develop the colors.

• The sugars will appear as pink or purple spots on the TLC plate.

Result:

• Observe the spots formed on the TLC plate after ninhydrin staining. Each distinct spot represents a different sugar component separated from the bark extract. The colors and positions of the spots indicate the presence and relative abundance of sugars.

Conclusion:

Thin-Layer Chromatography successfully separates the sugar components present in the bark extract. The sugars were visualized as pink or purple spots after treatment with ninhydrin solution, confirming the presence of carbohydrates in the extract.

Module 4: Biochemical Analysis  

4.1 Qualitative Tests for plant Metabolites (Protein, Carbohydrate, Alkaloids, Tannins)

Aim: To perform qualitative tests for the detection of proteins, carbohydrates, alkaloids, and tannins in plant extracts.

 Requirements:

Chemicals:

·         40% NaOH solution, 1% Copper sulfate solution, 0.2% Ninhydrin reagent, Fehling’s solution (A and B), Benedict’s solution, Wagner’s reagent, 1% Lead acetate solution, 5% Ferric chloride solution, 20% Sulfuric acid solution, Aqueous sodium hydroxide solution etc.

Apparatus:

• Test tubes, Test tube rack, Water bath, Pipettes, Droppers, Measuring cylinder, Heating source, Glass rods,  TLC plates (optional for advanced analysis) etc.

Plant Extracts:

• Plant extract rich in proteins, carbohydrates, alkaloids, and tannins (e.g., bark extract)

Introduction

Plant metabolites such as proteins, carbohydrates, alkaloids, and tannins play crucial roles in various physiological processes. This protocol aims to detect these metabolites in plant extracts through qualitative biochemical tests. These tests are based on chemical reactions that produce characteristic color changes or precipitates, indicating the presence of specific metabolites.

Principle: Each test is based on a specific biochemical reaction:

• Proteins react with copper sulfate in alkaline conditions (Biuret Test) or ninhydrin to form colored complexes.

• Carbohydrates reduce copper ions in Fehling’s and Benedict’s solutions to form colored precipitates.

• Alkaloids react with Wagner’s reagent to form precipitates.

• Tannins react with lead acetate, ferric chloride, or sodium hydroxide to give distinct colors or precipitates.

1. Test for Proteins

Biuret Test:

• Procedure:

  1. Take 2-5 ml of the plant extract.

  2. Add an equal volume of 40% NaOH solution.

  3. Add 2 drops of 1% copper sulfate solution.

  4. Observation: Violet color indicates the presence of proteins.

Ninhydrin Test:

• Procedure:

  1. Take 2-5 ml of the extract.

  2. Add 2 drops of 0.2% ninhydrin reagent.

  3. Gently heat the mixture.

  4. Observation: Pink or purple color indicates the presence of proteins or peptides.

2. Test for Carbohydrates

Fehling’s Test:

• Procedure:

  1. Take 2-5 ml of the plant extract.

  2. Add 5 mL of Fehling’s solution.

  3. Heat in a water bath.

  4. Observation: Yellow or red precipitate indicates the presence of reducing sugars.

Benedict’s Test:

• Procedure:

  1. Take 2-5 ml of the extract.

  2. Add 5 mL of Benedict’s solution.

  3. Heat in a water bath.

  4. Observation: Red, yellow, or green precipitate indicates the presence of reducing sugars.

3. Test for Alkaloids

Wagner’s Test:

• Procedure:

  1. Take 5-10 ml of the plant extract.

  2. Add a few drops of Wagner’s reagent.

  3. Observation: A reddish-brown precipitate indicates the presence of alkaloids.

4. Test for Tannins

Lead Acetate Test:

• Procedure:

  1. Take 5-10 ml of the plant extract.

  2. Add 0.5 mL of 1% lead acetate solution.

  3. Observation: A precipitate indicates the presence of tannins.

Ferric Chloride Test:

• Procedure:

  1. Take 5-6 ml of the plant extract.

  2. Add 0.5 mL of 5% ferric chloride solution.

  3. Observation: A dark bluish-black color indicates the presence of tannins.

Sodium Hydroxide Test:

• Procedure:

  1. Dissolve 5-6 ml of the extract in 0.5 mL of 20% sulfuric acid solution.

  2. Add a few drops of aqueous sodium hydroxide solution.

  3. Observation: A blue color indicates the presence of phenols.

Results:

• Proteins: Presence confirmed by violet color in the Biuret Test or pink/purple color in the Ninhydrin Test.

• Carbohydrates: Presence confirmed by yellow/red precipitate in Fehling’s Test or red/yellow/green precipitate in Benedict’s Test.

• Alkaloids: Presence confirmed by a reddish-brown precipitate in Wagner’s Test.

• Tannins: Presence confirmed by precipitate formation in the Lead Acetate Test, dark bluish-black color in the Ferric Chloride Test, or blue color in the Sodium Hydroxide Test.

Conclusion: This practical outlines qualitative biochemical tests to detect proteins, carbohydrates, alkaloids, and tannins in plant extracts. Each test relies on specific color changes or precipitate formation that confirms the presence of these metabolites, offering valuable insights into the biochemical composition of plant materials.

4.2: Study and Verification of Beer and Lamert’s Law

 AIM: To study and verify Lambert-beer’s law

Theory: The Beer-Lambert law explains that the amount of light absorbed by a solution is directly related to both the concentration of the absorbing substance and the path length through which the light travels. This means that, when using UV/Vis spectroscopy, the concentration of a substance in a solution can be determined if the path length remains constant. As the concentration of the colored solution increases, the amount of light absorbed also increases, while the amount of light transmitted decreases.

log ( I o / I t ) = A= ε c l

 Where

I o and I t are the incident and transmitted intensities,

A = absorbance and

ε is a constant i.e. absorptivity (formerly called the extinction coefficient ). If the concentration is measured in molL−1, the absorptivity is called the molar absorptivity .

A= ε c l

At constant length

A α c

 Requirements

·         Colorimeter, cuvette, six test tubes

·         Two Burettes or graduated cylinders two 100 mL beakers

·         0.01M KMnO4 solution

·         Distilled water, test tube rack, stirring rod, tissues (preferably lint-free)

Procedure:

(a) Determination of λmax
(b) Measuring Absorbance of Solutions at Different Concentrations at λmax

(a) Calculating λ max:
This value can either be referenced from tables of molar extinction coefficients or determined more precisely through a calibration curve.

1. Prepare a stock solution by dissolving 0.01M KMnO₄ (Molecular weight: 158.03 g/mol) in 100 mL of water and place it in a burette.

2. Turn on the instrument and computer, allowing a 30-minute warm-up period.

3. Set the instrument to display either % transmittance or absorbance, selecting the desired wavelength range.

4. Use a clean, dry glass or quartz cuvette (depending on wavelength) with a path length of 1 cm. Prepare a blank by filling the cuvette ¾ full with distilled water.

Note: Proper Handling of Cuvettes

• Wipe the outside of the cuvette with a lint-free tissue.

• Hold cuvettes by the top edges to avoid fingerprints.

• Remove air bubbles by tapping the cuvette gently on a hard surface.

• Ensure the light passes through the clear sides of the cuvette.

5. Using the blank cuvette (filled with distilled water), calibrate the colorimeter (set absorbance to 0, transmittance to 100%) using filter 1. Remove the blank cuvette afterward.

6. Fill a different cuvette with the KMnO₄ stock solution and record the absorbance reading in the table.

7. Repeat steps 5 and 6 for each filter across the wavelength range.

8. From the recorded data, identify λmax, the wavelength at which the solution shows the highest absorbance or optical density.

(b) Measuring absorbance of different concentration solutions

1.  Obtain small volumes 0.01M KMnO4 (Molecular weight 158.03 gm/mol) of solution and distilled water in separate beakers, fill in the separate graduated burettes

2.  Label five clean, dry, test tubes 1–5. Use Burettes to prepare five standard solutions according to the chart below. Thoroughly mix each solution with a stirring rod. Clean and dry the stirring rod between uses.

 Concentration can be calculate by

M1V1 = M2V2

You are now ready to collect absorbance-concentration data for the five standard solutions.

1.  Switch on the computer and/or the instrument powers; wait for 30 minutes for ‘warm-up’ of the instrument.

2.  In the instrument one can select light sources (UV and visible), choose the slit width, scan speed and % transmittance or absorbance display, wavelength range of interest, etc.

3.  Take two clean and dry glass (only for visible range scan) or quartz cuvettes with a given path length (say, 1 cm). Prepare a blank by filling a cuvette 3/4 full with distilled water and the other cuvette with aqueous KMnO4 solution with lowest concentration.

4.  Read the absorbance value displayed in the meter. When the displayed absorbance value stabilizes, record its value in your data table.

5.  Repeat the procedure for Test Tubes 2 to 5.Similarly spectral runs are done for all the other samples starting from the lowest concentrations to next higher concentrations of KMnO4. Every time one should rinse the cuvette taking a small portion of the solution to be analyzed next.

6.  Plot a curve between Absorbance v/s concentrations. Check whether it is a liner plot or not.

Result: A linear curve is obtained between Absorbance v/s concentrations that prove the existence of Lambert-Beer law.

Precautions: One should note that the Beer–Lambert law is obeyed by many substances mainly at low to moderate concentrations; therefore, dilute concentrations of the absorbing species should be measured. In practice it is advisable to measure absorbances in the range 0.1< A <1.0. Care must be taken to avoid any kind of chemical associations/dissociations of the absorbing species.

Module 5: Basic Molecular Technique

5.1 Genomic DNA Extraction:

Objective:
To extract high-quality genomic DNA from plant tissues using both conventional methods and commercially available kits.

Principle: Good quality DNA is essential for DNA manipulation experiments. The DNA extraction process involves the disruption of cell walls, membranes, and nuclear membranes to release DNA into a solution, while simultaneously removing proteins, polysaccharides, lipids, phenols, and other contaminants. The primary steps include cell disruption, DNA precipitation, and purification. Various components such as detergents, reducing agents, chelating agents, buffers, and salts facilitate this process, ensuring the extracted DNA is suitable for further molecular experiments.

Components of DNA Extraction:

1. Extraction Buffer:

   • Contains detergent like cetyl trimethyl ammonium bromide (CTAB) or SDS to break down membranes.

   • Reducing agent: β-mercaptoethanol aids in protein denaturation by breaking disulfide bonds.

   • Chelating agent: EDTA removes magnesium ions necessary for DNase activity.

   • Buffer: Usually Tris (pH 8) maintains pH levels.

   • Salt (NaCl): Neutralizes DNA charges, facilitating its precipitation.

2. Phenol-Chloroform Extraction:

   • Removes protein contaminants by mixing the nucleic acid solution with phenol, chloroform, and isoamyl alcohol. Proteins accumulate in the organic phase while nucleic acids remain in the aqueous phase.

3. Nucleic Acid Precipitation:

   • Alcohol precipitation (ethanol or isopropanol) is used to precipitate nucleic acids. Salt such as sodium acetate aids the process.

4. DNA Resuspension:

   • After precipitation, the DNA pellet is resuspended in sterile distilled water or TE buffer.

5. DNA Purification:

   • RNase A is used to remove RNA, followed by reprecipitation and phenol-chloroform extraction to purify the DNA.

A: Conventional DNA Extraction Protocol:

1. Reagents:

   • CTAB extraction buffer: 1.4 M NaCl, 100 mM Tris (pH 8.0), 20 mM EDTA, 2% CTAB, β-mercaptoethanol.

   • Isopropanol, Phenol (pH 8.0), Chloroform: Isoamyl alcohol (24:1).

   • TE Buffer (10mM Tris, 1mM EDTA).

   • RNase A (10 mg/ml) prepared in 10mM Tris-Cl, 15mM NaCl.

   • 70% Ethanol.

2. Materials:

   • Mortar and pestle, sterile glassware, centrifuge tubes, pipettes, and tips.

3. Procedure:

   • Weigh 2 g of young leaf tissue and grind it in liquid nitrogen.

   • Transfer to a 50 ml centrifuge tube containing 10 ml pre-warmed extraction buffer (65°C). Vortex and incubate for 1 hour.

   • Add 10 ml chloroform: isoamyl alcohol, mix by swirling, and centrifuge at 10,000 rpm for 10 minutes.

   • Transfer the aqueous phase to a new tube, add chilled isopropanol, and let DNA precipitate at -20°C for 30 minutes.

   • Spool out DNA, wash with 70% ethanol, and dry.

   • Resuspend DNA in TE buffer and treat with RNase A for 1 hour at 37°C.

   • Perform phenol-chloroform extraction and precipitate DNA using sodium acetate and ethanol.

   • Resuspend the DNA pellet in minimal TE buffer and store at -20°C.

B: DNA Isolation Using Kits:

1. Anion-Exchange Chromatography:

   • Utilizes the charged nature of nucleic acids to separate them from proteins and other contaminants.

   • The DNA binds to a column in low-salt conditions, followed by washing and elution using high-salt buffer.

2. Silica-Gel Membrane Technology:

   • DNA binds to silica in the presence of chaotropic salts, removing proteins and polysaccharides. DNA is washed and eluted in low-salt buffer for use.

Precautions:

• Immediately transfer ground plant tissue into extraction buffer to prevent degradation.

• Handle the phases carefully during phenol-chloroform extraction to avoid contamination.

• Minimize vigorous mixing to prevent DNA shearing.

• Avoid over-drying the DNA pellet to ensure easy resuspension.

• Use decontaminated materials to prevent contamination.

Results:

• The extracted DNA should appear as a clear, high-molecular-weight band on an agarose gel.

• The purity of DNA can be assessed through spectrophotometry, with an A260/A280 ratio of ~1.8 indicating good quality DNA.

• Yield and purity are dependent on the tissue type and method used, with modern kits often yielding higher quality DNA than traditional methods.

References:

1. Dellaporta, S.L., Wood, J., & Hicks, J.B. (1983). A plant DNA mini preparation: Version II. Plant Molecular Biology Reporter 1: 19-21.

2. Saghai-Maroof, M.A., Soliman, K.M., Jorgensen, R.A., & Allard, R.W. (1984). Ribosomal DNA spacer length polymorphism in barley. Proc. Natl. Acad. Sci. USA 81: 8014-8018.

Module 6. Soil Analysis Techniques

6.1 Determination of temperature and pH of different Soil Samples

Aims: To measure the temperature and pH of different soil samples to assess their suitability for agricultural and environmental purposes.

Objectives:

1. Determine soil temperature:
   Measure soil temperature using a Vernier temperature probe and analyze the effects of water addition and environmental changes (indoor vs. outdoor conditions).

2. Measure soil pH:
   Calibrate a pH meter and test soil samples to assess their acidity or alkalinity.

Materials Required:

o   Vernier temperature probe: To measure soil temperature.

o   LabQuest device: For data collection from Vernier temperature probe.

o   Graduated cylinder (25 ml): To measure water for soil sample.

o   pH meter: To measure soil pH.

o   Beakers (50 ml): For preparing soil and buffer solutions.

o   Glass rod: For stirring soil sample.

o   Buffer solutions (pH 4, 7, and 9.2): For pH calibration.

o   Distilled water: For preparing soil suspension.

Part 1: Determination of Soil Temperature

Procedure:

1. Sample Preparation:

   • Collect a cup of soil from the field and transfer it to a beaker. Leave the sample overnight in the classroom or greenhouse for testing the next day.

2. Set up the Equipment:

   • Connect the Vernier temperature probe to the LabQuest device.

   • Set the device to record temperature once per minute for 40 minutes.

3. Data Collection:

   • Place the probe into the soil sample and begin data collection when instructed.

   • After collecting 10 data points, add 25 ml of water to the soil sample using the graduated cylinder.

   • At the 20th minute, move the sample outside for 10 minutes.

   • At the 30th minute, move the sample back inside for the remaining 10 minutes.

4. Data Export and Cleanup:

   • After the 40th minute, stop the data collection and export the data into a spreadsheet for analysis.

   • Clean up the laboratory area and the utensils used.

5. Analysis of Data:

   • Analyze the temperature data, including:

     • Mean, median, mode, range, and standard deviation.

     • Maximum and minimum temperature values.

   • Determine whether the temperature sensor is accurate based on statistical analysis.

Part 2: Determination of Soil pH

Theory:
Soil pH is a measure of the hydrogen ion concentration (H⁺) in the soil solution and reflects the soil's acidity, neutrality, or alkalinity. It significantly affects nutrient availability to plants and microbial populations. The ideal pH range for most crops is between 5.5 and 6.5.

Procedure:

1. Calibrate the pH Meter:

   • Use buffer solutions with pH values of 4, 7, and 9.2 to calibrate the pH meter.

   • Insert the electrode alternately in beakers with the buffer solutions and adjust the pH as required.

2. Prepare Soil Sample for pH Testing:

   • Weigh 10.0 g of the soil sample and transfer it into a 50 ml beaker.

   • Add 25 ml of distilled water to the soil.

   • Stir the mixture thoroughly with a glass rod for 10 seconds.

3. Measure Soil pH:

   • Insert the electrode into the soil-water mixture.

   • Record the pH value displayed on the calibrated pH meter.

4. Observation:

   • Compare the results with the pH reaction ratings provided in the table below.

Analysis:

• Mean Temperature: ________

• Minimum Temperature: ________

• Maximum Temperature: ________

Conclusion and Result:

1. Soil Temperature:

   • Based on the data, the soil temperature ranged between ________ and ________°C. The average temperature was calculated to be ________°C, showing variations when water was added and when samples were moved outside and inside. These fluctuations suggest the probe's accuracy and the impact of environmental changes on soil temperature.

2. Soil pH:

   • The soil samples exhibited pH values ranging from ________ to ________. Based on the soil reaction rating, the samples were classified as __________ (e.g., neutral, slightly alkaline, etc.). This data can help determine soil amendments, such as the need for liming acidic soils or treating alkaline soils with gypsum for better crop productivity.

Module 7: Computer Techniques   

7.1. Study of photo micrographic techniques.

Aim:
To capture high-quality photomicrographs using the photomicrography technique with the help of a smartphone camera.

Introduction:

Photomicrography involves capturing images through a microscope. These images, known as photomicrographs, are valuable for botanists, researchers, and students to document and present microscopic findings. Traditionally, expensive photomicroscopes were required for this task. However, advancements in smartphone technology have made it possible to achieve similar results with the help of smartphone cameras and attachments. This practical protocol outlines two approaches: the traditional method using microscopes with mounted cameras, and the alternative method using smartphones with image editing applications.

Equipment:

• Microscope

• Smartphone with camera

Procedure:

Step 1: Prepare the Microscope

1. Clean the lenses: Ensure that both the objective lenses and the eyepiece are free of dust and smudges.

2. Set up illumination: Adjust the light source for even and adequate illumination of the sample.

3. Prepare the microscope for photography:

   • Image an empty field with transmitted light.

   • Turn off any gain or offset settings.

   • Adjust the camera’s exposure to show a neutral grey field.

   • Perform a white balance to ensure accurate color representation.

Step 2: Prepare the Slides

1. Clean the slides: Ensure both sides of the slide and cover glass are clean.

2. Set the slide on the stage: Place the specimen side facing the objective lens.

3. Select the field: Choose an appropriate field of view that includes the object you want to photograph.

4. High-magnification imaging: For highly magnified images, use a high-magnification objective lens, not the eyepiece.

5. Capture scale reference: It is recommended to photograph the scales of an objective micrometer to enable accurate size determination of objects in the final print.

Step 3: Smartphone Camera Protocol

1. Clean the smartphone lens: Dust and debris can degrade image quality. Clean the lens carefully to avoid flare and distortion.

2. Align the camera: Hold the smartphone camera steady and parallel to the surface of the microscope eyepiece for the best results.

3. Avoid mixed lighting: Use a single light source for neutral images. Avoid combining daylight, fluorescent, and tungsten lights.

4. Do not use the camera's flash: Use natural or microscope light instead. The camera’s flash is generally not powerful enough for high-quality photomicrographs.

5. Steady the camera: Holding the camera steady is crucial for sharp images. Mastering this technique may take some practice but is essential for successful smartphone photomicrography.

Results:

• Capture and save photomicrographic images using your smartphone camera (Flash off, ISO-Automatic, White Balance-Automatic).

• Paste the captured photomicrographs into your journal for documentation.

Conclusion:

By following the steps above, we will be able to produce clear, high-quality photomicrographs using both traditional microscopy techniques and smartphone technology.

7.2. Use of computer for preparation of Tables, Graphs and Presentations (MS Office)

Aim:

To familiarize students with basic computer skills using Microsoft Office for creating tables, graphs, and presentations.

Objectives:

1. Learn to create and format tables using MS Word and MS Excel.

2. Create graphs and charts in MS Excel for data visualization.

3. Design simple presentations using MS PowerPoint.

Materials Required:

• Computer with Microsoft Office (MS Word, MS Excel, and MS PowerPoint)

• Data for table and graph creation

1. Preparation of Tables in MS Word or Excel

Step 1: Open MS Word or Excel

• Open MS Word or Excel by clicking on the icon in your computer’s start menu or desktop.

Step 2: Inserting a Table in MS Word

• In MS Word: Go to the "Insert" tab, then click on "Table." Choose the number of rows and columns you want.

• In MS Excel: Cells are organized in rows and columns by default. You can start by selecting the cells to be part of your table or insert a new table using the "Insert" tab.

Step 3: Formatting the Table

• Adjust Columns/Rows: Hover over the edges of the cells to drag and adjust the row/column size as needed.

• Format Table: Use the “Table Design” or “Format” tab to apply various styles, like colors, borders, and shading to the table.

Step 4: Adding Data

• Click on the cells and start typing the required data into your table. Ensure data is aligned correctly for easy reading.

2. Creating Graphs in MS Excel

Step 1: Enter Data for the Graph

• Open Excel and enter your data into the cells. Ensure your data is well-organized, with labels for rows and columns.

Step 2: Select the Data

• Highlight the cells containing the data you wish to include in your graph.

Step 3: Insert Graph

• Go to the "Insert" tab, and in the “Charts” section, choose the type of graph you need (Bar Chart, Line Chart, Pie Chart, etc.).

Step 4: Customize the Graph

• Once the graph is inserted, you can customize it by clicking on the chart. Use the "Chart Design" and "Format" tabs to add titles, change colors, adjust axis labels, and more.

Step 5: Adjusting the Chart Type

• You can change the chart type by right-clicking on the chart and selecting "Change Chart Type."

3. Creating Presentations in MS PowerPoint

Step 1: Open MS PowerPoint

• Open PowerPoint and create a new presentation by clicking on "New" in the "File" tab.

Step 2: Create a New Slide

• Click on the “Home” tab, then “New Slide” to add a blank slide or choose from the available layouts.

Step 3: Add Title and Content

• Click on the text boxes to add a title and the content for your slide. You can customize the font, size, and color using the options in the “Home” tab.

Step 4: Insert Graphs or Tables

• To add tables or graphs, go to the "Insert" tab. Click on "Table" to insert a table or “Chart” to insert a graph.

• Select the type of graph you need, and a chart will appear. You can then edit the chart by adding your data.

Step 5: Add Visuals (Images, Icons)

• To insert pictures or images, click on the "Insert" tab and select "Pictures" to upload images from your computer.

• You can also use the "Icons" button to insert free icons into your presentation for visual enhancement.

Step 6: Design and Animations

• To make your presentation visually appealing, use the “Design” tab to apply a theme.

• Add transitions and animations between slides by selecting the “Animations” or “Transitions” tab.

Conclusion

By following these steps, basic-level we will be able to effectively create tables, graphs, and presentations using MS Office. It covers essential skills required for projects of basic research reports, and visual presentations.