FST 60293 Food Biochemistry

Observing the differences between plant cells (such as onion skin cells) and animal cells (such as cheek epithelial cells) under a microscope can be a fascinating way to understand cellular structures. Staining enhances the visibility of these structures, making it easier to distinguish between plant and animal cells. Here’s a guide on how to proceed with observation after staining:

Materials Needed

1. Microscope slides and coverslips

2. Microscope with at least 400x magnification

3. Stains:

   • Iodine solution (commonly used for plant cells)

   • Methylene blue (commonly used for animal cells)

4. Onion bulb (for plant cells)

5. Sterile cotton swab (for cheek cells)

6. Dropper and water for wet mounts

Procedure

1. Preparing the Onion Skin (Plant Cell) Slide

• Peel a thin, translucent layer of onion skin.

• Place it on a microscope slide.

• Add a drop of iodine solution; this will help stain the cell wall, nucleus, and sometimes starch granules.

• Carefully place a coverslip on top to avoid air bubbles.

2. Preparing the Cheek Epithelial (Animal Cell) Slide

• Gently scrape the inside of your cheek with a sterile cotton swab.

• Smear the collected cells on a microscope slide.

• Add a drop of methylene blue; it stains the nucleus and cytoplasm for better visibility.

• Place a coverslip over the sample carefully to avoid air bubbles.

3. Observing the Cells

• Start with a low power objective lens (10x) to locate the cells on both slides.

• Once located, switch to a higher magnification (40x or 100x) for detailed observation.

• Adjust the fine focus knob to get a clear view of cell structures.

Key Differences to Observe

1. Cell Wall (Plant Cells Only)

• In the onion cells, look for a rigid, rectangular cell wall surrounding each cell. The iodine will make this structure more visible.

• In the cheek cells, you will not see a cell wall, as animal cells only have a cell membrane.

2. Shape

• Onion cells will appear more regular and rectangular due to the cell wall, allowing them to fit together like bricks.

• Cheek cells will appear more irregular and rounded, with flexible shapes due to the lack of a cell wall.

3. Chloroplasts (Absent in Onion and Animal Cells)

• Although chloroplasts are common in green plant cells, onion cells do not contain chloroplasts since they are from an underground part of the plant.

• Cheek cells also lack chloroplasts.

4. Central Vacuole (Plant Cells)

• In onion cells, you may notice a large central vacuole occupying much of the cell’s interior. This vacuole often appears as an empty space due to the cell's storage of water.

• Cheek cells lack a large central vacuole; animal cells, if they have vacuoles, are much smaller.

5. Nucleus

• Both types of cells will have a clearly visible nucleus after staining, though it may be more prominent in cheek cells with methylene blue.

• The position of the nucleus in plant cells is often pushed to the edge due to the large vacuole, while in cheek cells, it may be more centrally located.

Isolating and observing chloroplasts from spinach or other green leaves is a straightforward process and provides a fascinating look at these vital organelles involved in photosynthesis. Here’s a step-by-step guide for isolating and viewing chloroplasts:

Materials Needed

1. Fresh spinach leaves (or other green leaves)

2. Blender or mortar and pestle

3. Cold isolation buffer (e.g., 0.35M sucrose solution or a buffer containing sucrose, Tris, and MgCl₂)

4. Filter (cheesecloth or a coffee filter)

5. Centrifuge (if available, a table-top centrifuge works best)

6. Microscope slides and coverslips

7. Microscope (400x magnification or higher)

8. Pipette or dropper

9. Beaker and test tubes

Procedure

1. Prepare the Spinach Leaf Suspension

• Rinse the spinach leaves with distilled water to remove any dust or impurities.

• Tear the leaves into small pieces and place them in a blender.

• Add a small amount of cold isolation buffer to help keep the chloroplasts intact and stable.

• Blend the leaves on low speed for about 10–20 seconds, or gently grind them using a mortar and pestle until you have a thick, green leaf slurry.

2. Filter the Leaf Slurry

• Pour the leaf slurry through a cheesecloth or coffee filter into a beaker to remove larger pieces of leaf tissue. This will result in a green liquid containing suspended chloroplasts and other cell components.

3. Separate the Chloroplasts

• Transfer the filtered solution to a test tube.

• If you have a centrifuge, place the test tube in it and centrifuge the solution at around 1,000-2,000g for about 5 minutes. The chloroplasts should form a green pellet at the bottom of the tube.

• Carefully pour off the supernatant (the liquid above the pellet), being careful not to disturb the chloroplast pellet at the bottom.

4. Resuspend the Chloroplasts

• Add a small amount of fresh isolation buffer to the tube and gently mix to resuspend the chloroplasts. This will help create a concentrated chloroplast suspension suitable for microscopic observation.

5. Preparing the Slide

• Using a pipette or dropper, place a small drop of the chloroplast suspension onto a clean microscope slide.

• Gently place a coverslip over the drop to avoid air bubbles.

6. Observing Chloroplasts Under the Microscope

• Place the slide under the microscope and start with a lower magnification (e.g., 10x) to locate the chloroplasts.

• Once located, switch to higher magnification (e.g., 40x or 100x) to observe their shape, structure, and movement.

Observing Chloroplasts

When observed under the microscope, chloroplasts from spinach or other green leaves will appear as small, oval-shaped structures with a green color due to chlorophyll. They may appear to move slightly in the buffer, a phenomenon known as cytoplasmic streaming if they are in intact cells.

You should be able to observe:

• Shape and Size: Chloroplasts are generally oval or disc-shaped and fairly uniform in size.

• Color: Their green color is due to chlorophyll pigments, which are critical for photosynthesis.

• Possible Movement: In some cases, intact chloroplasts in plant cells may exhibit a slow, flowing motion within the cell, which is called cytoplasmic streaming.

This process will provide a clear view of chloroplasts and highlight their role in the cell.

Qualitative Tests for Carbohydrates

Objective:
To identify the presence of different types of carbohydrates (monosaccharides, disaccharides, and polysaccharides) using standard biochemical tests.

Materials Needed:

• Test tubes

• Benedict's solution

• Iodine solution

• Molisch's reagent

• Barfoed's reagent

• Glucose, sucrose, and starch solutions (5%)

• Distilled water

• Pipettes

• Hot water bath

Procedure:

1. Molisch Test (General Test for Carbohydrates):

   • Add 2 mL of each carbohydrate solution into separate test tubes.

   • Add 2 drops of Molisch's reagent to each.

   • Slowly add 1 mL of concentrated sulfuric acid along the wall of the test tube.

   • Observe for a purple ring at the interface.

2. Benedict's Test (Reducing Sugars):

   • Add 2 mL of Benedict's solution to 2 mL of each carbohydrate solution.

   • Heat in a boiling water bath for 5 minutes.

   • Observe the color change (green, yellow, orange, or red indicates reducing sugar).

3. Iodine Test (Starch):

   • Add 2-3 drops of iodine solution to each test tube containing the carbohydrate solution.

   • Observe the color change (blue-black indicates the presence of starch).

4. Barfoed's Test (Monosaccharides):

   • Add 2 mL of Barfoed's reagent to 2 mL of each carbohydrate solution.

   • Heat in a boiling water bath for 3 minutes.

   • Observe for red precipitate (indicates monosaccharides).

Observations and Results:
Record the results for each test and classify the carbohydrate type.

Estimation of Reducing Sugars by Fehling’s Method

Objective:
To quantify reducing sugars in a given sample using Fehling's solution.

Materials Needed:

• Fehling’s solution A and B

• Glucose solution (standard)

• Test sample solution

• Burette

• Pipette

• Conical flask

• Hot water bath

Procedure:

1. Prepare standard glucose solutions of varying concentrations (e.g., 0.2%, 0.4%, 0.6%, 0.8%, and 1%).

2. In a conical flask, mix 5 mL each of Fehling's solution A and B.

3. Add 10 mL of the glucose solution to the Fehling's mixture.

4. Heat the mixture in a boiling water bath until a brick-red precipitate forms.

5. Repeat the procedure with the test sample solution.

6. Record the volume of glucose solution required to completely reduce the Fehling’s reagent.

Calculation:
Use the standard graph of glucose concentration vs. volume of Fehling's reagent reduced to determine the concentration of reducing sugar in the test sample.

Observations and Results:
Compare the volume of the sample required with the standard curve to estimate the reducing sugar concentration.

Qualitative Tests for Proteins

Objective:
To identify the presence of proteins and distinguish between different types using biochemical tests.

Materials Needed:

• Test tubes

• Pipettes

• Biuret reagent

• Ninhydrin solution

• Xanthoproteic reagent (concentrated nitric acid)

• Albumin solution (5%)

• Distilled water

Procedure:

1. Biuret Test (General Test for Proteins):

   • Add 2 mL of albumin solution to a test tube.

   • Add 2 mL of Biuret reagent and mix.

   • Observe for a violet or purple color, indicating the presence of proteins.

2. Ninhydrin Test (Detection of Free Amino Acids):

   • Add 2 mL of albumin solution to a test tube.

   • Add 2 mL of Ninhydrin solution.

   • Heat the mixture in a water bath for 5 minutes.

   • Observe for a blue or purple color, indicating free amino acids.

3. Xanthoproteic Test (Detection of Aromatic Amino Acids):

   • Add 2 mL of albumin solution to a test tube.

   • Add a few drops of concentrated nitric acid.

   • Heat gently, then cool and neutralize with sodium hydroxide.

   • Observe for a yellow color turning orange, indicating aromatic amino acids.

Observations and Results:
Record the color changes for each test to confirm protein presence and composition.

Estimation of Protein by Lowry's Method

Objective:
To estimate the protein concentration in a sample using Lowry’s method.

Materials Needed:

• Protein standard solution (e.g., bovine serum albumin, BSA)

• Lowry reagent (Folin-Ciocalteu phenol reagent, sodium carbonate solution, copper sulfate, and sodium tartrate)

• Test sample

• Spectrophotometer

• Pipettes and cuvettes

Procedure:

1. Prepare standard protein solutions of varying concentrations (e.g., 20 µg/mL, 40 µg/mL, etc.).

2. Take 1 mL of each standard solution or test sample in separate test tubes.

3. Add 5 mL of the prepared Lowry reagent to each test tube and mix.

4. Incubate the mixture at room temperature for 10 minutes.

5. Add 0.5 mL of Folin-Ciocalteu reagent to each tube and mix immediately.

6. Incubate at room temperature for 30 minutes.

7. Measure the absorbance at 660 nm using a spectrophotometer.

8. Plot a standard curve using the protein concentrations and their absorbance values.

9. Determine the protein concentration in the test sample using the standard curve.

Observations and Results:
Use the standard curve to estimate the protein concentration in the test sample and compare it with expected values.

Qualitative test for lipids

Objective:
To identify the presence of lipids using standard qualitative tests.

Materials Needed:

• Test tubes

• Sudan III solution

• Ethanol

• Water

• Lipid sample (e.g., oil, butter)

• Filter paper

Procedure:

1. Sudan III Test:

   • Add 2 mL of lipid sample to a test tube.

   • Add a few drops of Sudan III solution and shake gently.

   • Observe for a red-stained oil layer, indicating the presence of lipids.

2. Solubility Test:

   • Add a small amount of lipid sample to separate test tubes containing water and ethanol.

   • Observe solubility; lipids are insoluble in water but soluble in ethanol.

3. Grease Spot Test:

   • Place a drop of the lipid sample on a piece of filter paper.

   • Allow it to dry and observe under light.

   • A translucent spot confirms the presence of lipids.

Observations and Results:
Record color changes, solubility patterns, and the appearance of translucent spots.

Saponification Reaction

Objective:
To demonstrate the saponification reaction of lipids to produce soap and glycerol.

Materials Needed:

• Test tubes

• Lipid sample (e.g., coconut oil or butter)

• Sodium hydroxide solution (NaOH)

• Ethanol

• Water bath

• Phenolphthalein indicator

Procedure:

1. Take 2 mL of the lipid sample in a test tube.

2. Add 2 mL of ethanol and mix well to dissolve the lipid.

3. Add 2 mL of NaOH solution.

4. Heat the mixture in a water bath for 10-15 minutes.

5. Allow the solution to cool and add a few drops of phenolphthalein.

6. Add distilled water to observe soap precipitation.

Observations and Results:
Note the formation of soap as a solid precipitate and confirm the reaction by the appearance of a pink color with phenolphthalein (indicating unreacted alkali).

Estimation of Lipids by Gravimetric Method

Objective:
To estimate the lipid content in a given food sample using the gravimetric method.

Materials Needed:

• Food sample (e.g., seeds, nuts)

• Mortar and pestle

• Soxhlet apparatus (or alternate extraction setup)

• Petroleum ether (solvent)

• Weighing balance

Procedure:

1. Weigh a known amount of the food sample (~5 g).

2. Grind the sample using a mortar and pestle.

3. Place the sample in the Soxhlet extractor and add petroleum ether as the solvent.

4. Heat the setup to allow lipid extraction for 4-6 hours.

5. After extraction, evaporate the solvent to leave behind lipids.

6. Weigh the extracted lipids.

Calculation:

Lipid Content (%)=(Weight of extracted lipidWeight of sample)×100\text{Lipid Content (\%)} = \left(\frac{\text{Weight of extracted lipid}}{\text{Weight of sample}}\right) \times 100Lipid Content (%)=(Weight of sampleWeight of extracted lipid​)×100

Observations and Results:
Record the weight of the lipid extracted and calculate the percentage of lipids in the sample.

Qualitative Analysis of Nucleic Acids

Objective:
To identify the presence of DNA and RNA in a sample using qualitative biochemical tests.

Materials Needed:

• Nucleic acid samples (DNA and RNA solutions)

• Diphenylamine reagent (for DNA test)

• Orcinol reagent (for RNA test)

• Dische’s reagent (for DNA detection)

• Test tubes

• Boiling water bath

• Pipettes

Procedure:

1. Diphenylamine Test for DNA

• Take 2 mL of the DNA sample in a test tube.

• Add 2 mL of diphenylamine reagent to the test tube.

• Heat the mixture in a boiling water bath for 10 minutes.

• Observe the development of a blue color, which indicates the presence of DNA.

2. Orcinol Test for RNA

• Take 2 mL of the RNA sample in a test tube.

• Add 2 mL of orcinol reagent to the test tube.

• Heat the mixture in a boiling water bath for 10 minutes.

• Observe the development of a green color, which indicates the presence of RNA.

3. Dische’s Test for DNA

• Take 2 mL of the DNA solution in a test tube.

• Add 2 mL of Dische’s reagent.

• Heat the mixture in a boiling water bath for 5-10 minutes.

• Observe the appearance of a blue-green color, which confirms the presence of deoxyribose in DNA.

Observations and Results:

• Note the color changes for each test.

• Record observations