13.2: Exercise
Exercise 1: Separation of Plant Pigments Using Chromatography
In this lab, you will first extract pigments from spinach leaves and then separate pigments from one another using a technique called chromatography. Chromatography is used to separate chemicals based on their varying solubilities in selected solvents. Recall from our discussions of the Chemistry of Life (textbook chapter), molecules and compounds are classified as polar or nonpolar , and that “like dissolves like.” In other words, polar compounds dissolve in polar solvents, but not in non-polar solvents, and vice versa. An example of this concept is seen when oil, a nonpolar substance, comes in contact with water, a polar substance. The two substances do not mix; they are repelled by one another. Chromatography works by separating chemicals according to their varying degrees of polarity. In chromatography experiments, there are two “phases,” the stationary phase, which does not move, and the mobile phase, which travels across the stationary phase.
In this experiment, you will use paper chromatography . Paper, which is made of cellulose, is very polar and acts as the stationary phase. The solvent you will use is a mixture of petroleum ether and absolute acetone, both of which are nonpolar; the solvent acts as the mobile phase. Extracted plant pigment is applied to the paper, and the paper is placed in the solvent. As the solvent moves up the paper through capillary action, it carries the pigments along with it. Pigments that are more polar are attracted to the polar cellulose molecules in the stationary phase (the paper), so they move more slowly as the solvent travels up the paper. Nonpolar pigments are more strongly attracted to the nonpolar solvent and tend to stay in solution longer, thus moving farther up the paper. The pigments are carried at different rates because they are not equally soluble. Since different pigment molecules have different molecular structures and varying degrees of polarity, this technique works well to separate pigments from one another, giving us a clear look at each pigment individually.
The distance that each pigment travels will be unique to that pigment based on the solvent selected. These migration distances can be used to calculate the ( retention factor(opens in new window) ) value, which is simply calculated by dividing the distance traveled by the pigment over the distance traveled by the solvent.
Materials
- Spinach leaves
- Electronic balance / weigh boats
- A small amount of clean quartz sand
- Mortar and pestle
- Acetone (keep this in the hood at all times)
- Test tube
- Chromatography paper (11 cm square)
- Metric ruler
- Chromatography jars - glass quart jars (which will hold 11 cm square paper) with lids or can cover with aluminum foil
- Pencil
- Glass capillary tubes
- Small metric rulers
- Chromatography solvent (Use 9 parts petroleum ether and 1 part acetone)
- Waste container for discarded acetone/chlorophyll extract
- Handheld UV light (Turn lights off in lab and shine handheld UV light on students’ chlorophyll extracts to cause them to fluoresce red)
Procedures
Pigment Extraction
- Obtain spinach leaves. Tear leaves into small pieces, discarding the large midvein. Weigh out approximately 4.0 grams of leaf tissue.
- Place leaf tissue plus a pinch of clean quartz sand into a clean mortar and pestle; grind to a fine pulp.
- Add 6mL of acetone to the pulp and continue to grind.
- When leaf tissue is thoroughly ground into a paste-like consistency, allow the solution to rest for 1 minute. Carefully pour off the liquid portion of the mixture into a clean test tube, leaving the pulpy remains in the mortar. *Note: Dispose of acetone/chlorophyll waste in the waste container in the fume hood. Wash the mortar and pestle well with warm soapy water, rinse, and set aside to dry.
- You may wish to examine your acetone/chlorophyll extract under UV light. Turn off the lights in the lab and shine the fluorescent black light onto the extract. Do not look into the light directly. The chlorophyll will fluoresce red.
- Can you explain why the extract containing the chlorophyll pigment turns from a green color to a fluorescent red color when exposed to UV light?
Pigment Separation
NOTE: The organic solvents used in this step are extremely volatile and flammable. The chromatography jars must be kept in the fume hood at all times.
Paper chromatography requires that the atmosphere within the chromatography jar be completely saturated with solvent. Be sure that the lid or aluminum foil covering the jar stays in place before and during chromatogram development.
- Prepare chromatography sheet: Use the pencil to draw a line 1.5 - 2 cm from the bottom of the paper. This line will serve as a guide when applying the spinach extract. It will help ensure that the extract is applied evenly, in a straight line, and at a level above the solvent in the chromatography jar.
- Apply pigment: Using a glass capillary tube, apply the pigment extract to the paper in a linear series of small dots (follow the line you created as a guide for application). NOTE: Leave a 1 cm margin at each edge that is free of pigment extract. DO NOT TOUCH THE PAPER; oils from your skin can interfere with the process.
- After you have completed the pigment line across the chromatography paper, allow the extract to dry for about 5 min. Go back and re-apply more pigment over the first line, allowing each application to dry before reapplying more extract. Repeat the procedure until all the pigment extract is used, or until you have applied enough extract (consult with your instructor). Allow the sample to dry.
- Once your sample has dried, take your chromatography paper to the fume hood. Add approximately 20 mL of chromatography solution to the jar (this may have already been done for you). This should result in the solvent being about 1 cm deep from the bottom of the jar. Roll the paper into a cylinder so that the line faces outward and is toward the bottom of the jar. You can secure this cylinder shape by using a small stapler to staple the sides together at the top and the bottom of the cylinder. Place the rolled paper into the jar so that it is in the chromatography solution. The chromatography solution should not be higher than the line on your chromatography paper.
- Cover the jar with the lid or aluminum foil and observe as the chromatography solvent (the mobile phase) travels up the paper (the stationary phase).
- When all of the pigments are clearly separated and before the solvent front has reached the top edge, remove the chromatogram and allow it to dry in the hood. Make sure to use a pencil to draw a line to indicate where the solvent front ended.
- You should be able to see at least four distinct bands. (There may be as many as six bands.)
- Make a sketch of your chromatogram using the template that follows. Using colored pencils, note the color of the bands. Use Table 1 to help you identify each pigment. On your sketch, label each band with the name of the pigment. Mark the distance (in cm) from the initial pigment band to each colored band as well as the total distance from the initial pigment band to the solvent front.
Sketch of Resulting Chromatogram
|
Pigment |
Band Color |
|---|---|
|
Beta-carotene |
Orange |
|
Xanthophylls |
Yellow |
|
Chlorophyll a |
Blue-green |
|
Chlorophyll b |
Yellow-green |
- Use the data from your sketch to complete the data table below.
- Calculate the value and include that in your data table.
|
A |
B |
C |
D |
E |
|---|---|---|---|---|
|
Name of Pigment |
Description of color |
Distance solvent front traveled from initial pigment band |
Distance pigment traveled from initial pigment band |
Rf value of pigment (column D / column C) |
Exercise 2: Analysis of the Absorption Spectrum of Leaves
While we know most leaves are green, we have now seen that other pigments are also found in the leaves. The green color that we observe is a consequence of reflected light in the wavelengths of about 550-500nm in size ( Fig. 3 ). In the autumn months of the year, many plants stop producing chlorophyll, revealing the colors of the other pigments beneath. Every chemical has a specific set of wavelengths it can absorb and thus be identified. By analyzing the absorption spectrum of leaf pigments, we can infer what wavelengths of light will be useful in photosynthesis.
Materials
- PASCO Wireless Spectrometer
- iPad or laptop to connect the spectrometers to
- 4 cuvettes
- Leaf samples
- 3 test tubes
- 100 mL 95% ethanol
- 20 mL graduated cylinder
- Scissors
- Mortar and pestle set up from the previous section
Procedure
- Connect the spectrometer to a Laptop via a USB cable or wirelessly to an iPad.
- Open the PASCO spectrometer application.
- Select the ANALYZE Solution setting from the menu at the top.
- Choose CALIBRATE DARK from the menu at the bottom. Cover the sample well with your finger to block light until a check mark appears indicating that calibration is complete.
- Fill a cuvette ¾ full with ethanol. Only handle the cuvettes from the ribbed sides since fingerprint smudges on the clear sides will affect light passing through. Place the cuvette in the well so that the clear side is facing the light source.
- Choose CALIBRATE REFERENCE from the menu until the checkmark appears. You do not need to cover the cuvette with your finger for this step.
SAMPLE PREPARATION:
- Obtain 3 different samples of plant leaves.
- Label 3 test tubes: Sample 1, Sample 2, Sample 3
- Tear the first sample of plant leaves into small pieces, discarding the large midvein. Weigh out approximately 4.0 grams of leaf tissue.
- Place the leaf tissue plus a pinch of clean quartz sand into a clean mortar and pestle; grind to a fine pulp.
- Add 6mL ethanol to the pulp and continue to grind. Make sure to use ethanol and NOT acetone. The cuvettes are plastic and acetone will cloud the plastic and not allow light to pass through freely.
- When the leaf tissue is thoroughly ground into a paste-like consistency, allow the solution to rest for 1 minute. Carefully pour off the liquid portion of the mixture into a clean test tube, labeled sample 1, leaving the pulpy remains in the mortar.
- Repeat the extraction for the remaining two leaf samples. Be sure to add the appropriate leaf extraction mixture to the correctly labeled test tubes.
DATA COLLECTION
- Place the first leaf sample extraction mixture into a cuvette in the well and press START RECORDING. The graph that appears is the absorbance spectrum of the extract. If any of the peaks are flattened, add more ethanol to the cuvette to further dilute the sample.
- When the peaks have stabilized, select STOP RECORDING and name your run for the sample analyzed.
- Use the ADD COORDINATE tool to find the peak of each sample and SNAPSHOT each of the sample runs. Record the values in a data table.
- Table 2 shows the absorption spectrum of some common plant pigments. Use this information to predict what pigments were in your samples.
|
Pigment |
Peak Absorption Wavelengths (nm) |
Sample 1 |
Sample 2 |
Sample 3 |
|---|---|---|---|---|
|
Chlorophyll A |
430 and 662 |
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Chlorophyll B |
453-642 |
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Carotenoids |
460-550 |
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Anthocyanins |
520* |
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Xanthophyll |
494 |
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Betalains |
535 or 480* |
*peaks may vary and are pH-dependent.
Exercise 3: Measuring the Rate of Photosynthesis
Materials:
- Five spinach leaves (Dark green, fresh spinach leaves produce better, faster results.)
- Two cups (Short clear plastic cups which hold ~300 mL work best.)
- Index cards
- Forceps
- Plastic petri dish
- 500 mL of water with 0.2% sodium bicarbonate (NaHCO3) (or baking soda; you may prefer to use 0.3%-0.5% sodium bicarbonate which will result in a faster rate of photosynthesis and more rapid floating for the leaf disks)
- 500 mL of water without sodium bicarbonate (NaHCO3)
- Two drops of dilute detergent (To prepare the dilute detergent, add approximately 5 mL of dishwashing liquid soap to 250 mL of water. Aliquot into microfuge tubes and label with a “DD”)
- Hole punch (or a straw)
- Two 10 ml syringes without needles (syringes can be rinsed, dried and reused)
- Lamp with 23 W spiral compact fluorescent bulb (If you have a clamp lamp, obviously this could be attached to a ring stand or other vertical pole; if you don't have enough vertical poles, you could rig up lamp support using a cardboard box.)
- Stopwatch
Employing Steps in the Scientific Method:
- Record the Question that is being investigated in this experiment. ________________________________________________________________
- Record a Hypothesis for the question stated above. ________________________________________________________________
- Predict the results of the experiment based on your hypothesis (if/then). ________________________________________________________________
- Perform the experiment below and collect your data.
Procedure:
- Label one cup 'sodium bicarbonate' and fill it about one-quarter full (~75 mL) with the sodium bicarbonate solution. Label the second cup 'water' and fill it about one-quarter full (~75mL) with water. Add one drop of dilute detergent (microfuge tube labeled “DD”) to each cup.
- Next, you will prepare your leaf disks, taking care not to damage them. First, prepare two index cards by folding each of them in half and then unfolding them and laying them flat. Use the hole punch to prepare the leaf disks. Punch out a piece of leaf tissue, avoiding large veins. (If you are using a hole punch, keep it clamped shut as you tap the punch on the table to release the leaf disk onto the piece of paper.) Repeat until you have 10 leaf disks on each index card.
- Remove the plunger from a syringe, and use one of the folded papers to pour 10 leaf disks into the syringe. Tap them down to the tip of the syringe.
- Replace the plunger, and push it down to about the 1 mL mark, being careful not to squash the leaf pieces.
- Suck up ~5 mL of the sodium bicarbonate/detergent solution into the syringe. Hold the syringe upright and push out as much of the air as possible.
- Put your thumb over the tip of the syringe, and pull back slowly on the plunger to about the 10 mL mark. Additionally, while pulling back on the plunger and keeping your thumb over the tip of the syringe, shake the solution in the syringe. This creates a vacuum and pulls air out of the leaf discs. Tilt and swirl the syringe to make sure all disks are submerged in the solution, then hold it for 10 seconds, and then gently let go of the plunger without removing your thumb from the tip of the syringe. The plunger will “snap” back into position and the solution will enter the leaf disks. If the leaf disks drop to the bottom of the solution in the syringe, you are done. If not, do this again.
- Remove the plunger and empty the disks into the cup containing sodium bicarbonate solution. Then swirl the cup to dislodge any disks that are stuck to the side of the cup. Make sure that all of the disks are settled on the bottom of the cup.
- Repeat steps 3 - 7 but use the water/detergent solution instead of sodium bicarbonate.
- Place both cups under a bright light. Place half a petri dish full of water on top of each cup to act as a temperature buffer.
Safety Precaution
Be careful to keep all liquids away from the light source and electrical cord.
- At the end of each minute, record the number of floating disks (any disk that is no longer touching the bottom) in the following table.
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Minute |
In Water/Detergent |
In Bicarbonate/Detergent |
|---|---|---|
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1 |
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2 |
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3 |
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4 |
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5 |
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6 |
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7 |
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8 |
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9 |
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10 |
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11 |
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12 |
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13 |
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14 |
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15 |
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16 |
Extension Activity: (Optional)
The results of this experiment can be presented graphically. The presentation of your data in a graph will assist you in interpreting your results. Based on your results, you can complete the final step of scientific investigation, in which you must be able to propose a logical argument that either allows you to support or reject your initial hypothesis.
- Graph your results using the data from Table 2, using different symbols for the leaf disks in water vs. the leaf disks in bicarbonate solution.
- What is the dependent variable? Which axis is used to graph this data? ______________________________________________________________________
- What is your independent variable? Which axis is used to graph this data? _____________________________________________________________________
Exercise 4: Observing and Quantifying Stomata (Optional)
Leaves are the primary food factories in a typical plant. Powered by the sun’s energy, leaves take in water and carbon dioxide and produce oxygen and glucose through photosynthesis . Leaves have specialized cells on their surfaces called guard cells which surround openings called stomata (singular = stoma). Stomata are tiny openings in the epidermis of terrestrial (land) plants. Gases, primarily O2 and CO2, can pass through these openings to allow for photosynthesis to occur. Likewise, H2O will evaporate out of the stomata, leading to potential dehydration.
Stomata typically remain open during the day (since sunlight stimulates them to open) and close at night, when photosynthesis doesn’t occur, allowing for water conservation. Stomata can also close during especially hot days, during droughts, or as a response to plant growth regulators. Desert plants have a special form of photosynthesis which allows them to open their stomata at night, to perform gas exchange, and close them during the day to conserve water.
Materials:
- Potted plant with leaves
- Bottles of clear nail polish
- Clear packing tape
- Microscopes
- Microscope slides
- Forceps
- Scissors
- Calculator
Employing Steps in the Scientific Method:
- Record the Question that is being investigated in this experiment. ________________________________________________________________
- Develop a hypothesis about the number of open stomata found on the upper side of a leaf as compared to the lower side of the leaf. ___________________________________________________________
- Develop a hypothesis about the number of open stomata found on the upper side of a leaf as compared to the lower side of the leaf. Write your hypothesis in the space below. ____________________________________________________________
- Predict the results of the experiment based on your hypothesis (if/then). ________________________________________________________________
- Perform the experiment below and collect your data.
Procedure:
- Obtain 2 cut sections of leaves from the plant provided in the lab.
- Using a small amount of clear nail polish, coat the underside (lower surface) of one of the leaves with a THIN layer. Set the leaf on a paper towel (painted surface up) and allow it to dry completely (~ 30 min.)
- Coat the upper surface of the remaining leaf in the same way.
- As the nail polish dries, it will conform to the surface of the leaf.
- Place a piece of the clear cellophane tape over the entire surface of the leaf covered with the nail polish. Use your fingertip to gently adhere the tape to the entire surface of the leaf, rubbing back and forth until you feel you have pressed the tape into the surface of the leaf.
- Use a corner of the tape to gently peel back the tape. While peeling back the tape you will start to separate the dried nail polish from the surface of the leaf. This is the leaf impression you will examine under the microscope.
- Tape each of your peeled impressions to a clean microscope slide. Use scissors to trim away any excess tape.
- Examine the leaf cell imprints under a total magnification of 40x. Scan the slide until you find a good area where you can see plenty of the stomata. *Note: You may need to adjust the iris diaphragm to reduce the amount of light and add contrast to the slide to more clearly see the stomata.
- Increase the total magnification to 100x for counting.
- Each stoma is surrounded by two sausage-shaped cells that are smaller than the surrounding epidermal cells. These cells are called guard cells and, unlike other cells in the epidermis, contain chloroplasts.
- Identify the epidermal guard cells and the stomata. Fix the field of your microscope in place, and count the number of visible stomata. Stomata on the edge of the field of view are to be counted! Record this number in Table 3 under “ View 1 ”.
- Move the field of view to a new position on the leaf and repeat the procedure. Record this number in Table 3 under “ View 2 ”.
- Do a third count in a new position and record this number in Table 3 under “ View 3 ”. (Do three different counts for the upper surface leaf imprint and the lower surface leaf imprint.)
|
Surface of Leaf |
# of Stomata: View 1 |
# of Stomata: View 2 |
# of Stomata: View 3 |
Average # of Stomata |
|---|---|---|---|---|
|
Upper |
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|
Lower |
Extension Activity: (Optional)
The results of this experiment can be presented graphically. The presentation of your data in a graph will assist you in interpreting your results. Based on your results, you can complete the final step of scientific investigation, in which you must be able to propose a logical argument that either allows you to support or reject your initial hypothesis.
- Graph your results using the data from Table 3.
- What is the dependent variable? Which axis is used to graph this data? ______________________________________________________________________
- What is your independent variable? Which axis is used to graph this data? _____________________________________________________________________