Materials Included In Kit

Additional Materials Required

Prelab Preparation

1. Slurry of each of the extract powders.

a. Pour each of the extract powders into separate, labeled 50-mL beakers.
b. Add 8 mL of acetone to each beaker.
c. Swirl for several minutes.
d. Cover to prevent evaporation of the acetone.

2. Prepare TLC plates for each lab group.

1. Cut the TLC sheets into plates approximately 6.5 cm x 3 cm.
2. Be careful not to scrape any of the silica gel from the plates. Note: Some silica gel will chip from the edges of each plate—this is not a problem.

Safety Precautions

Acetone and the chromatography solvent are flammable and dangerous fire risks; toxic by ingestion and inhalation. This lab should be performed only in an operating chemical fume hood or well-ventilated area. Wear chemical splash goggles, chemical-resistant gloves and a chemical-resistant apron. Remind students to wash hands thoroughly with soap and water before leaving the laboratory. Please consult current Safety Data Sheets for additional safety, handling and disposal information.

Disposal

Please consult your current Flinn Scientific Catalog/Reference Manual for general guidelines and specific procedures, and review all federal, state and local regulations that may apply, before proceeding. Acetone, chromatography solvent, spinach extract and blue-green algae extract may be disposed of according to Flinn Suggested Disposal Method #18a. TLC plates may be disposed of according to Flinn Suggested Disposal Method #26a.

Lab Hints

• Enough materials are provided in this kit for 30 students working in pairs or for 15 groups of students. This laboratory activity can reasonably be completed in one 50-minute class period. The pre-laboratory assignment may be completed before coming to lab, and the data compilation and calculations may be completed the day after the lab.
• Good technique is important to achieving clean separations in thin-layer chromatography. Common sources of student error include “overloading” the TLC plates by placing too much extract on the initial spot and band broadening that occurs because the initial spot is too large. Another common problem is “staining” that occurs when the pigment spots are submerged below the solvent level.
• Allowing enough time for the development of the TLC is critical. The plate must be left in the development beaker long enough for the solvent to be drawn up near the top of the plate. Do not stop the development until the solvent front nears the top of the plate. Underdevelopment will lead to incomplete separation. Typical time for a 6.5 cm TLC plate is 10 minutes.
• The chromatography solvent contains 80% petroleum ether and 20% acetone. Do not discard leftover chromatography solvent—the solvent may be recycled. Save it for use by another class or for another chromatography experiment. Do not leave the chromatography solvent uncovered for long periods of time—one component may evaporate faster than the other, changing the polarity of the mixture.

Teacher Tips

• Many plant pigments fluoresce under ultraviolet light. Once students’ TLC plates have dried, shine a UV light source very close to the plate—you should see many or all of the pigments fluoresce! You may even find pigments that were not visible under room light. Note: A black-light bulb will not work.

• Have students prepare extracts from other plants. Simply soak leaves from various plants in acetone to extract pigments. They may need to tear and/or grind the leaves (with a mortar and pestle and a small amount of clean sand) to extract the pigments. Additional chemicals and materials are available from Flinn Scientific, Inc.

Answers to Prelab Questions

1. Read the Background section and the Procedure—why is a pencil rather than a pen used to mark the “starting line” in step 3?
   

   Pen ink may contain pigments that will dissolve in the solvent and interfere with the separation.
   

2. The plant extracts in this experiment will be drawn up the capillary tube and the solvent will flow upward through the TLC plate, each by capillary action. What is capillary action and what causes it?
   

   Capillary action refers to the spontaneous rise of a liquid in small tubes or fibers, or to the wetting of a solid (e.g., paper or fabric) in contact with a liquid. Capillary action is responsible for the rise of sap in plant fibers and the flow of blood through capillaries. Capillary action is due to attractive forces (adhesion) between the molecules of the liquid and the walls of the vessel.
   

3. What is the solvent front and how is its migration on the TLC plate measured?
   

   The solvent front is the leading edge of the solvent as it travels up the TLC plate. The distance the solvent front migrates is measured from the pencil line where the plant extract is spotted to the maximum height the solvent rose.

Sample Data

Spinach

Blue-green algae

Answers to Questions

1. Compare and contrast the pigments observed on the spinach TLC plate with the pigments on the blue-green algae TLC plate.
   

   Both samples exhibited chlorophyll a. Both also contained a xanthophyll, although they appeared to be different pigments. The blue-green algae contained phycobilins. The spinach contained carotenoids and chlorophyll b.

2. What factors are involved in the separation of the pigments?
   

   The polarity of the pigments varies because of differences in the structures of the pigment molecules. The different polarities cause the pigments to adsorb differently on the TLC plates. The polarity of the solvent and the affinity for the adsorptive surface affect the separation of the pigments. The time and distance allowed for the separation to occur also affect the separation of the pigments.
   

3. Refer to step 8 in the Procedure: Why was it necessary to keep the chromatography solvent level in the beakers below the sample extracts spotted on the TLC plates?
   

   If the chromatography solvent is above the extract spots on the TLC plates, some of the extract will dissolve directly in the solvent and will not travel as a distinct band on the TLC plates.
   

4. Which pigment directly captures light energy? What are the roles of the other pigments?
   

   Chlorophyll a is the only pigment that is able to directly capture energy from light and convert it to chemical energy. The other pigments expand the range of wavelengths of light absorbed, but that energy must be transferred to chlorophyll a.

References

Bregman, A. A. Laboratory Investigations in Cell Biology, 2nd ed.; John Wiley & Sons: New York, 1987; pp 119–123.

Bold, H. C.; Wynne, M. J. Introduction to the Algae, 2nd ed.; Prentice-Hall: Englewood Cliffs, NJ, 1985, p 39.

Green, N. P. O.; Stout, G. W.; Taylor, D. J. Biological Science: Organisms, Energy, and Environment: 2nd ed.; Saper, R., Ed.; Cambridge University: Cambridge, MA, 1990; pp 255–257.

Russo, T.; Meszaros, M. W. Vial Organic; Flinn Scientific: Batavia, IL, 1996; pp 25–33.

Wilkins, M. B., Ed. Advanced Plant Physiology; Pitman: Marshfield, MA, 1984; pp 221–224.

Introduction

Chromatography is a popular method used to separate organic compounds for identification or purification. Discover for yourself the wide variety of different pigments found in spinach and blue-green algae by analyzing a chromatograph.

Concepts

• Chromatography

• Plant pigments
• Photosynthesis

Background

I. Photosynthetic Pigments

A pigment is a molecule or compound that absorbs certain wavelengths of visible light very strongly and transmits or reflects the remaining wavelengths of visible light. Different pigments have different molecular structures and thus reflect different specific wavelengths of visible light. The characteristic color of an organism is due to the reflection of specific wavelengths of light by pigments within the organism (see Table 1). The major pigments of photosynthetic organisms are the chlorophylls. There are two types of green chlorophyll, called chlorophyll a and chlorophyll b. Chlorophyll a and chlorophyll b are found in all plants, most algae and some bacteria. Chlorophylls c and d are not green and only occur in some algae and bacteria.

In addition to the chlorophylls, autotrophs also contain other pigments used to collect light energy (see Figure 1). These other pigments are known as accessory pigments. There are more than 1000 known accessory pigments. The accessory pigments may be divided into two classes: the carotenoids and the phycobilins. The phycobilins are red to blue in color and only occur in Cyanobacteria and Rhodophyta. In Table 1, three major phycobilins are listed; they are allophycocyanin, phycocyanin and phycoerythin. The carotenoids include carotenes and xanthophylls. The xanthophylls are yellow to red in color and occur in plants, algae, bacteria and diatoms. In Table 1, lutein and the pigments ending in -anthin are xanthophylls. The carotenes, including lycopene, are yellow, orange or red in color and occur in plants, algae, bacteria and diatoms. The colors of the accessory pigments are masked in plants during the summer by the high concentration of chlorophyll a. In autumn, the decrease in the amount of daylight causes the plant to break down the chlorophyll, which allows the brilliant reds, oranges and yellows of the accessory pigments to become visible in the fall leaves. Of all of the pigments, only chlorophyll a directly captures light energy and converts it to chemical energy. Chlorophyll b and the accessory pigments assist the process by expanding the range of wavelengths absorbed (see Figure 1) and then transmitting the energy to chlorophyll a.

The differences in the molecular structure of various pigments mean that the pigments all have different polarities as well. Variations in polarity make it possible to separate plant pigments using a process known as chromatography. There are many different types of chromatography, but most work on the principle of adsorbtion. An adsorbent is a solid which is capable of attracting and binding the components in a mixture (see Figure 2). In this laboratory, a thin layer of silica spread onto a thin plate of non-reactive plastic will act as the adsorbent. Many other chemicals may also be used as the adsorbent in thin-layer chromatography (TLC). Other examples of chromatography used by scientists to separate molecules based upon the concept of adsorbtion are paper chromatography and gas chromatography.

In this laboratory, a suspension of plant pigments will be “spotted” onto the surface of a silica TLC plate and a solvent will then allowed to seep or flow through the silica on the surface of the TLC plate separating the different pigments. The separation occurs because one of the components in the mixture is more strongly adsorbed onto the silica than another. As a result it will spend a smaller fraction of time free in solution and will move up the TLC plate more slowly than the solvent. Components that are not strongly adsorbed onto the silica will spend a greater fraction of time free in solution and will move up the TLC plate at a faster rate. This “partitioning” of the components of a mixture between the silica and the solvent separates the components and gives rise to different color bands that become visible on the surface of the TLC plate.

How far each pigment migrates depends upon many factors, including how high the solvent is allowed to rise, the type of adsorbent, the type and concentration of the solvent, the temperature of the experiment and the distance of the starting point from the pool of solvent. In order to compare values, scientists calculate the relative mobility of each pigment. Scientists report the experimental conditions and the relative mobility, called the retention fraction (R_f), for each pigment and compare their results with those from other scientists conducting similar experiments. R_f is defined as the fractional rise of the pigment compared to the rise of the solvent (see Equation 1).

Experiment Overview

The purpose of this activity is to separate plant pigments by thin-layer chromatography. The retention factor, R_f, will be calculated for each pigment.

Materials

Prelab Questions

1. Read the Background section and the Procedure—why is a pencil rather than a pen used to mark the “starting line” in step 3?
2. The plant extracts in this experiment will be drawn up the capillary tube and the solvent will flow upward through the TLC plate, each by capillary action. What is capillary action and what causes it?
3. What is the solvent front and how is its migration on the TLC plate measured?

Safety Precautions

The plant extract slurry contains acetone. Acetone and the chromatography solvent are flammable and dangerous fire risks. They are also toxic by ingestion and inhalation. This lab should be performed only in an operating chemical fume hood or well-ventilated area. Wear chemical splash goggles, chemical-resistant gloves and a chemical-resistant apron. Wash hands thoroughly with soap and water before leaving the laboratory.

Procedure

1. Label two 50-mL beakers with your group name or number. Label one beaker “spinach extract” and the other “blue-green algae extract.”
2. Touching only the edges of the TLC sheet, use scissors to cut the TLC sheet into two small plates approxi-mately 6.5 cm x 1.5 cm. Be careful not to scrape any of the silica gel off the plates—this will adversely affect results. Note: Some silica gel may chip from the edges of each plate—this is not a problem.
3. Using a pencil, draw a faint line 0.5 cm from the bottom edge of each TLC plate (see Figure 3).
4. You are now ready to spot the TLC plate using the capillary tubes. Dip the capillary tube into the slurry of one of the plant extracts. The solution will be drawn up the tube. Remove the capillary and about 1 cm of solution should remain in the tube due to capillary action. Briefly and gently touch the tip of the capillary tube to the center of the pencil line on the TLC plate, keeping your index finger over the end of the tube, so that only a small amount of solution is transferred with each touch. It is important to keep the spot as small as possible. Let the solvent evaporate before touching the capillary tube to the plate again. Blow gently on the spot to help it evaporate. Touch the capillary to the same spot again. Remove the capillary and gently blow on the spot to evaporate the solvent.
5. Repeat step 4 six to ten times or until a rather dark spot is present on the TLC plate. The spot should be approximately 5–6 mm in diameter when completed. Note: Too much extract on the initial spot will cause the resulting pigment bands to broaden and tail. Too little extract will create very faint bands that are difficult to analyze.

6. Repeat steps 4–5 for the second plant pigment extract.
7. Use a Beryl-type pipet or medicine dropper to transfer 2 mL of chromatography solvent into the bottom of each 50-mL beaker.
8. Carefully place each TLC plate, sample end down, into the appropriate 50-mL beaker. Note: The upper level of the solvent must not touch the sample extract (see Figure 4). If it appears that the sample will be within the solvent, take out the sample before the TLC plate touches the solvent and remove some of the chromatography solvent from the beaker.
9. Carefully cover each beaker with a watch glass or Parafilm to minimize evaporation of the solvent. Note: Do not disturb the TLC plate.
10. When the solvent front (top of the solvent) has moved to within 0.5–1 cm from the top of the TLC plate, remove the plate from the 50-mL beaker. Replace the watch glass on top of the 50-mL beaker to prevent the solvent from evaporating.
11. Use a pencil to immediately mark the location of the solvent front before it evaporates. Measure the distance from the pencil line to the solvent front in millimeters. Record this distance on the Plant Pigment Chromatography Worksheet.
12. Mark the center of each pigment band with a pencil.
13. Measure the distance each pigment migrated from the sample spot to the center of each separated pigment band. Record the distance, in millimeters, that each pigment moved on the worksheet.
14. Calculate the R_f value for each pigment and record this value for each pigment on the worksheet.
15. Refer to Table 1 in the Background section to identify each pigment band. Record the identity of each band on the worksheet.
16. Consult your instructor for appropriate disposal procedures.

Student Worksheet PDF

10117_Student1.pdf