In the Laboratory
Student Understanding of Chromatography: A Hands-On Approach Robert Curtright Lincoln Northeast High School, 2635 North 63rd St., Lincoln, NE 68507 Randy Emry Lincoln Southeast High School, 2930 South 37th St., Lincoln, NE 68506 John Markwell* Department of Biochemistry, University of Nebraska, Lincoln, NE 68588-0664
Research scientists depend upon using the right tools. The development and understanding of the “tools of the trade” go hand in hand with progress in science endeavors. Chromatography in all its forms is an essential tool for scientists and a common practice in chemistry, biology, and biochemistry. The principles of chromatography are essentially the same for paper chromatography, thin-layer chromatography (TLC), and high-pressure liquid chromatography (HPLC). Chemistry students armed with an understanding of the principles of polarity and chemical partitioning that underlie these research tools will undoubtedly be better prepared for original scientific investigation. For this reason we developed a series of student exercises intended to develop a better understanding of chromatography. The exercises employ plant pigments, which our students seem to find interesting and relevant. A collaborative approach leads students to a level of competence that permits them to complete an open-ended exercise. Background Many beginning chemistry students already have some experience with the technique of chromatography before they take chemistry. Their prior experiments may have been to separate the constituent pigments in a plant leaf (1) or an ink. Some have reported innovative and inexpensive ways of separating plant pigments (2, 3) . The primary objective of these exercises was often to demonstrate the fractionation of a chemical mixture into its components. The student usually does this exercise or similar ones without much opportunity to gain a broad-based understanding of the principles of chromatography. Our objective was to revisit this common exercise to provide students with an understanding of the chemical principles involved. Several types of plant material may be suitable for this exercise. Red- or purple-leafed plants are a likely source of both anthocyanins and chlorophylls. Plants with variegated leaves may or may not have both anthocyanins and chlorophyll, but they are a reasonable material for investigation. From a local plant nursery we obtained a list of common plants that possess anthocyanins throughout the growing season, such as purple-leaf plum, red-leaf barberry, and redvein prayer plant. Because of its colorful variegated leaves, we selected the Coleus plant as a familiar and visually striking example of the diversity of plant pigments. The pigments responsible for this array of color in a Coleus leaf represent *Corresponding author. Email:
[email protected].
molecules having a wide range of polarities. These pigments, which serve in light-harvesting, photochemical transduction, and UV protection, are of general interest to biologists and chemists (4) and of critical importance to plants (5). In the exercises described here, students will observe a demonstration resulting in the partitioning of pigments from a multicolored Coleus leaf. The exercises provide both an opportunity for students to learn how the polarity of the pigment molecules determines interaction with selected solvents and an involved discussion on the whole procedure. As the exercises continue, students use this understanding to develop a system that will separate the different pigments using a relatively inexpensive thin-layer chromatography (TLC) system. The solvents used in this activity are (in order of increasing polarity) 2-propanol, acetone, methanol, and water. It should be noted that the type of polarity normally used to explain chromatography is different from the concept of polarity generally used in organic chemistry (6 ). In the latter, polarity is generally equated with the strength of the dipole moment. In terms of chromatography, benzene is considered to be a more polar solvent than cyclohexane, although neither has a dipole moment. Methanol is used to release chlorophyll, carotenoids, and anthocyanins from plant leaves because of its intermediate polarity. Students must mix different proportions of water with the more nonpolar molecules of acetone and 2-propanol to produce a satisfactory developing solution for chromatography. Chemical Safety CAUTION: Instructors should be aware that acetone, methanol, and 2-propanol are flammable solvents. All have flammability ratings of 3 on their material safety data sheets (MSDS). Store these solvents in a cool, dry, well-ventilated flammable-liquid storage area. In terms of potential health hazards, methanol is of greater concern than the other two solvents. Methanol has an MSDS health rating of 3, whereas the ratings for both acetone and 2-propanol are 1. If a chemical fume hood is not available for these exercises, consideration should be given to adequate ventilation. Care should be taken to dispose of solutions containing these solvents in accordance with pertinent regulatory guidelines. Demonstration and Discussion We suggest that an initial demonstration period be used to introduce the concept of polarity and partitioning and to let students see how pigments are extracted from leaves by
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grinding in methanol. For the demonstration, plant pigments can be extracted from Coleus leaves using a mortar and pestle with methanol as the solvent. Solid plant debris is removed from the resultant extract with a tabletop centrifuge or by filtration through filter paper. Once the pigments are separated from the debris, their partitioning can be demonstrated by preparing a bilayer of cooking oil and vinegar. The oil and vinegar do not mix owing to the fact that the strong attraction between water molecules excludes the oil molecules into a separate layer (7). The free energy for attraction between oil molecules is approximately equal to the free energy for attraction between oil and water molecules. The free energy for self-attraction of water molecules is more than three times greater (8), which drives the thermodynamic preference for minimization of water–oil contacts. Partitioning of plant pigments is achieved by adding the cleared extract sample into a test tube containing the bilayer of oil and vinegar. The contents of the test tube are mixed and allowed to stand until layers can be observed. Separation of the oil and vinegar layers after mixing can be expedited by use of a tabletop centrifuge. After separation the nonpolar oil layer will have collected nonpolar chlorophyll and carotenoid pigment molecules and the lower vinegar layer will have collected the more polar anthocyanin pigment molecules. The acidic environment of the vinegar enables visualization of the anthocyanins in this layer, as the color of this class of pigments is pH-sensitive (9). Following the demonstration with the vinegar and oil, which should be familiar to the students, we repeat the partitioning of the leaf extract with a vinegar and hexane system. This introduces the concept of using organic solvents and the rapidity of the separation creates a favorable impression. A fume hood should be employed for all but microscale amounts of hexane. The instructor should again explain that pigments partition or distribute into either the oil or the vinegar layer as a function of the polarity of the pigment molecule. It can be further explained that different molecules will have different partition coefficients for two-phase systems and will differentially distribute between the two phases. As a practical example resulting from a knowledge of partitioning, it can be explained that a grass stain on clothes will not wash out with just water because the chlorophyll in the stain is not soluble in water. To help students comprehend partitioning, we provide the following metaphor as a post-demonstration discussion (10). (It may be more appropriate to use a different metaphor with your own students.) On the day of a football game there is a large gathering for music lovers at opposite sides of the football stadium on campus. One of the two gathering sites provides country–western music entertainment; at the other a band is playing rock music. People tend to collect at the end of the stadium where the music is more liked (or less disliked). The result is a separation or partitioning of the crowd. We explain that just as the two bands at opposite ends of the stadium reflect different musical tastes and seldom play the same music, the solvents in common salad dressing, a mixture of vinegar and oil, or in the hexane and vinegar mixture, will not mix with each other and partition into two separate phases. Many molecules, if added to this bilayer system, will tend to distribute differentially on the basis of their affinity for a polar (vinegar is aqueous and mostly water) or a nonpolar (oil or hexane) environment. Thus anthocyanins added to a bilayer of oil and vinegar will partition 250
into the vinegar phase because they are polar molecules and their ability to hydrogen bond with the polar water molecules makes them soluble in this environment. Chlorophyll will partition into the oil because it is nonpolar and will not be accommodated by the water molecules. The next step in the desired cognitive development is to explain that chromatography is, in a sense, similar to a two-phase system (11). One difference, however, is that one phase is a mobile solvent and the other is static and composed of a solid adsorbent. Molecules subjected to chromatography are continually subjected to a differential distribution ratio between the stationary adsorbent and the mobile solvent, and thus can be fractionated by differential mobility. A great advantage of this technique is that the two systems need not be immiscible. This permits great flexibility in manipulation of the polarity of the mobile solvent phase. Student Exercise After the demonstration described above and the ensuing discussion, students are challenged to concoct a solvent system that will cause both the chlorophyll and the anthocyanin molecules to migrate during the TLC chromatography and to clearly separate from one another. They are to adhere to the following guidelines: 1. Use a three-component mixture of 2-propanol, water, and 10% acetone. 2. All the pigments must migrate from the origin. 3. The goal is to separate the chlorophylls from the anthocyanins. 4. Separation of carotenoids is of secondary importance, but separation of all three classes of molecules (anthocyanins, chlorophylls, and carotenoids) would indicate a superior accomplishment.
The polar anthocyanin and nonpolar chlorophyll molecules are soluble in methanol, a solvent of intermediate polarity. Using techniques observed in the demonstration, students use methanol to extract pigments from Coleus leaves. Methanol (2.5 mL) is added to 0.5 g of Coleus leaf and ground in a mortar with pestle to release pigments. The resultant extract is centrifuged for several minutes in a clinical centrifuge or filtered to remove solid plant debris. The developing solution (or mobile phase) for the chromatography is a mixture of PAW (2-propanol–acetone– water). The students’ task is to develop the proportions of a mixture of PAW that will be capable of both moving and separating chlorophyll and anthocyanins. To limit the number of possible solutions, the students are to keep the acetone concentration constant at 10%. Thus mixtures could have compositions such as PAW 45:10:45, or PAW 5:10:85, or PAW 37:10:53. The numbers refer to the volume (in mL) of each component used to make the final solvent. The reagents in PAW mix in a nonadditive fashion, so each one needs to be measured separately. To conserve chemicals the PAW mixtures were usually made to produce 10-mL volumes (e.g., PAW 4.5:1.0:4.5). Plastic-backed Silica Gel 60 plates are cut with scissors to have dimensions of about 3 × 8 cm. The Coleus extracts are spotted onto the plates with microcapillary tubes. The total volume of pigment necessary for visual monitoring of the fractionation is determined by visual inspection. The TLC plate is developed in a 250-mL beaker containing enough
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Figure 1. Contrasting chromatograms depicting pigment migration at the two extremes of solvent polarity. The numbers at the top of the figure represent the composition of the PAW developing solvent. (This figure appears in color on page 148.)
migration of pigments was not substantially different from that in the PAW 3:1:6 mixture. Examination of this series of plates (Fig. 2) shows that the anthocyanins begin to migrate from the origin with solvents more polar than an 8:1:1 mixture and move with the solvent front with solvents more polar than a 5:1:4 mixture. Chlorophyll starts to move off the origin with solvents less polar than a 4:1:5 mixture and migrates with solvents in a 6:1:3 or less polar mixture. In fact, the PAW 6:1:3 supports maximum migration of all the pigments. Tracing either anthocyanin or chlorophyll migration from plate to plate provides a pattern of migration of both pigments similar to a graph of chromatographic mobility versus mobile solvent polarity. In nearly all the plates, a yellow band, carotenoid pigments, can be observed. As is desirable for collaborative studies, there will be no one “best” solvent. Groups of students may argue for the superiority of different systems on the basis of individual criteria. An Opportunity for Student-Designed Exercises
Figure 2. An array of chromatograms developed in PAW solvent that increases in polarity from left to right. The PAW composition is indicated by the numbers above each chromatogram. (This figure appears in color on page 148.)
PAW mixture to cover the bottom. The beaker is covered with plastic wrap to contain solvent fumes. The plates are developed for 15–20 minutes, which allows enough time for migration of pigments in the mobile phase. Making their own developing solution gives students a chance to discover the principles of chromatography. For example, an examination of the extremes and midrange of PAW mixtures illustrates how well pigment molecules migrate in each of the PAW solvents (Fig. 1). In the PAW 9:1:0 solvent the anthocyanins failed to migrate off the origin. This solvent is apparently too nonpolar to dissolve the anthocyanins. At the other extreme, PAW 0:1:9, the chlorophyll was apparently too nonpolar to migrate in the polar solvent provided. Each plate had the same run time. The solvent fronts corresponded closely to the leading pigment. The mobile phase (solvent) migrates more rapidly in the PAW 9:1:0 than in any of the others. The middle plate permits both movement and some separation of the pigments. We have found that the learning atmosphere seems to be stimulated when students work in small collaborative groups. Additionally, this procedure is so rapid that students are able to attempt several combinations of PAW in the same lab period. Students measured the success of each experiment by evaluation against the suggested guidelines above. Results and Discussion An entire array of chromatograms can be produced by a collaborating group of students in a fairly short time. Figure 2 shows chromatograms developed in various PAW mixtures ranging from PAW 9:1:0 to PAW 3:1:6. PAW mixtures having less 2-propanol and more water are not pictured because the
Once the students have found a suitable developing solution, they should be encouraged to use TLC as a tool for new experiments. For example, one group of advanced chemistry students at Northeast High School (Lincoln, NE) used TLC to monitor the production of anthocyanins during the month of September. The burning bush (Euonymus alata) will have anthocyanins present even in early summer if the leaves are exposed to direct sunlight. The bush used in this experiment was in a shaded area and did not have detectable amounts of anthocyanins in early September. By the last week of September, however, the majority of students found evidence of anthocyanin production. Perhaps there is a local plant that appears green until autumn when the leaves turn scarlet, orange, or red. Students could study this plant for pigmentation changes. Are the anthocyanins always present and simply masked by chlorophyll, or are they produced in response to the changing autumn conditions? Sugar maple (Acer saccharum) and sumac (Rhus glabra or Rhus typhina) are two plants that might merit students’ investigation. In the winter perhaps students could investigate anthocyanins in poinsettia (Euphorbia pulcherrima) to learn more about anthocyanin production in these plants. During the spring months, the newly unfurled leaves of some trees such as cottonwood (Populus deltoides) have a noticeable tinge of red, and as the leaves mature this color is seemingly lost. A study could be conducted to learn if anthocyanin production is halted or if anthocyanin synthesis continues but these red pigments are masked by the green chlorophyll pigments. Conclusion Students’ development and understanding of the “tools of the trade” requires time and experience. Students need more than just an introduction to chromatography to use this technique as a tool permitting them to ask and answer their own questions. Chromatography is commonly used in recipe-like applications with little or no emphasis as how it works. Learning the principles behind chromatography has value as a science lesson in itself. For this reason we believe that hands-on education illustrating the principles involved in
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common techniques such as chromatography is an important ingredient in science education. Acknowledgments We are grateful to Jon Wright for assistance with this exercise. Development of the exercise was funded by grant DUE-9550791 from the National Science Foundation. This is manuscript No. 981-2, College of Agricultural Sciences and Natural Resources, University of Nebraska. Literature Cited 1. Kimbrough, D. R. J. Chem. Educ. 1992, 69, 987.
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2. 3. 4. 5. 6. 7. 8. 9. 10. 11.
MeWald, W.; Rodolph, D.; Sady, M. J. Chem. Educ. 1985, 62, 530. Lalitha, N. J. Chem. Educ. 1994, 71, 432. Diehl-Jones, S. M. J. Chem. Educ. 1984, 61, 454. Harborne, J. B. Comparative Biochemistry of the Flavonoids; Academic: New York, 1967. Touchstone, J. C.; Dobbins, M. F. Practice of Thin Layer Chromatography; Wiley: New York, 1983. Pravia, K.; Maynard, D. F. J. Chem. Educ. 1996, 73, 497. Tanford, C. The Hydrophobic Effect: Formation of Micelles and Biological Membranes; Wiley: New York, 1980. Curtright, R. D.; Rynearson, J. A.; Markwell, J. J. Chem. Educ. 1994, 71, 682. Starkey, R. J. Chem. Educ. 1986, 63, 514. Brimley, R. C.; Barrett, F. C. Practical Chromatography; Reinhold: New York, 1953.
Journal of Chemical Education • Vol. 76 No. 2 February 1999 • JChemEd.chem.wisc.edu
