Spinach

Sandra K. Koster, Ph.D. Lecturer
Department of Chemistry
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Isolation of Chlorophyll and Carotenoid Pigments from Spinach

Adapted from: Pavia, D.L.; Lampman, G.M.; Kriz, G.S.; Engel, R.G. Introduction to Organic Laboratory Techniques: A Microscale Approach 3^rd Edition Saunders College Publishing: New York, NY, l999 and also Quach, H. T.; Steeper, R. L.; Griffin, G. W., J. Chem. Educ., 2004, 81, 385-387.

The leaves of plants contain a number of colored pigments generally falling into two categories, chlorophylls and carotenoids. Chlorophylls a and b are the pigments that make plants look green. These highly conjugated compounds capture the (nongreen) light energy used in photosynthesis.

Carotenoids are part of a larger collection of plant derived compounds called terpenes. These naturally occurring compounds contain 10, 15, 20, 25, 30 and 40 carbon atoms which suggest that there is a compound with five carbon atoms that serves as their building block. Their structures are consistent with the assumption that they were made by joining together isoprene units, usually in a "head to tail" fashion. Isoprene is the common name for 2-methyl-1,3-butadiene. The branched end is the "head" and the unbranched is the "tail". That isoprene units are linked in a head to tail fashion to form terpenes is known as the isoprene rule. Carotenoids are tetraterpenes (eight isoprene units). Lycopene, the compound responsible for the red coloring of tomatoes and watermelon, and b -carotene, the compound that causes carrots and apricots to be orange, are examples of carotenoids.

b -Carotene is also the coloring agent used in margarine. When ingested b -Carotene is cleaved to form two molecules of vitamin A and is the major dietary source of the vitamin. Vitamin A, also called retinol, plays an important role in vision.

Spinach leaves contain chlorophyll a and b and b -carotene as major pigments as well as smaller amounts of other pigments such as xanthophylls which are oxidized versions of carotenes and pheophytins which look like chlorophyll except that the magnesium ion Mg⁺⁺ has been replaced by two hydrogen ions 2H⁺. In this experiment we will isolate and separate the spinach pigments using differences in polarity to effect the separation. Since the different components are colored differently, we can follow this separation visually. The structures of the major components are given below. Notice that since b -carotene is a hydrocarbon it is very nonpolar. Both chlorophylls contain C--O and C--N bonds which are polar and also contain magnesium bonded to nitrogen which is such a polar bond it is almost ionic. Both chlorophylls are much more polar than b -carotene. If you look carefully you can see that the two chlorophylls differ only in one spot. Chlorophyll a has a methyl group (--CH₃) in a position where chlorophyll b has an aldehyde (--CHO). This makes chlorophyll b slightly more polar than chlorophyll a. After we isolate the pigment mixture from the leaves in a hexane solution we will use the difference in polarity to separate the various pigments using column chromatography. We will analyze the original extract and the pigment fractions using thin layer chromatography, which also separates based on polarity.

Chlorophyll a

Blue-green, polar

C₅₅H₇₂MgN₄O₅

M. W. 893.5026

Chlorophyll b

Green, polar

C₅₅H₇₀MgN₄O₆

M. W. 907.4862

beta-Carotene--yellow, nonpolar

C₄₀H₅₆

M.W. 536.8824

PROCEDURES

Isolation of pigment from leaves:

<>Weigh about 1.0 g of fresh spinach leaves (or other fresh green leaves; avoid using stems or thick veins). Place them in a mortar along with 1.0 g of anhydrous magnesium sulfate and 2.0 g fo sand. Grind with a pestle until a light green power is obtained (about 5-10 minutes). Transfer the mixture to a centrifuge tube. Add 2.0 ml of acetone. Rinse the mortar and pestle with another 2.0 ml of acetone, and transfer the remaining mixture to the centrifuge tube. Cap and shake the mixture and then allow it to stand for 10 minutes. Stir or shake and let stand an additional 5 minutes. Using a Pasteur pipette, or by careful pouring, transfer the liquid to a vial. Rinse the solid with 1.0 ml acetone and transfer this liquid to the same vial. Label the vial E for extract. If time is short, label the vial with your name and store it open in your desk until the next laboratory period. If there is time to continue the experiment the same day, use a gentle stream of air or nitrogen in the hood to effect a quicker evaporation. If this technique is used, add 1 ml of hexane to the dried vial and if water appears as a lower layer, pipette just the hexane into a new vial.

Column Chromatography

Column chromatography involves the separation of compounds by the same mechanism as other chromatographic techniques, i.e. differences in partitioning between mobile and stationary phases. (See "Chromatography" in Pavia). It is like the other methods in that a stationary phase is placed in a support through which the mobile phase is passed. The stationary phase serves as an adsorbent. Many compounds with varying functional groups may be used as the stationary phase and several types of interactions can aid in developing the desired separation (i.e. hydrogen bonding, electrostatic interactions, Van der Waals forces, etc.) The major advantages of column chromatography are its ability to handle large amounts of material and the ability to change the eluting solvent throughout the course of the elution. This allows one to remove one component while a desired product remains essentially unmoved. Then a solvent change moves the desired product through the column. Solvent changes may include such things as changes in polarity, changes in pH, or changes in ionic strength. The last two are used largely in biological separations. Thus, by varying the stationary phase and by changing solvent or solvent systems, an efficient separation may be achieved. In this experiment the florisil separates components primarily on the basis of polarity; the more polar components are held to the florisil more tightly and therefore move through the column more slowly. Increasing the polarity of the solvent moves all components faster but has the largest effect on polar compounds.

Column Chromatography of Spinach Pigments:

A clean, dry Pasteur pipette is aligned vertically (very important), a very small plug of glass wool is pushed to the bottom of the column and a thin layer of about 1/2 cm of sand is added. Silica gel (enough to fill pipette about 2/3 full) is added and gently tapped to level it. Your instructor may instruct you to use florisil or alumina in place of the silica gel. One half cm of sand is added carefully to the top of the column. Again, tap gently to achieve a flat surface. If the column is to be stored for use later, it may be covered with parafilm to protect the packing material from moisture. The following solvents should be obtained and brought to your work space before you proceed: 5 ml hexane, 5 ml 90% hexane-10% acetone solution (by volume), 10 ml 70% hexane-30% acetone solution, 5 ml acetone. Since our chromatography column is really a Pasteur pipette we don't have a stopcock at the bottom to stop the flow of solvent during the procedure and so all materials needed should be at the bench before the chromatography is started. Also have several vials labeled 1, 2, 3, etc., as well as a beaker to hold waste chromatography solvent.

<>If your extract vial has dried out, reconstitute it by adding about 1/2 ml of hexane. Set aside about 1/4 of your extract (from vial E) to use for later thin layer chromatography. Once the procedure is started, it should not be stopped: the silica gel must be kept wet with solvent all the time. When you are ready to begin the column chromatography, place the waste solvent beaker under the Pasteur pipette and add about 2.0 ml of hexane to the top of the pipette. As the silica gel is wetted, the hexane will flow into the beaker. As soon as the solvent is drained so that the top layer of sand is just covered by the solvent, start adding your spinach extract to the top of the column. As the extract drains onto the florisil or silica gel, the pigments will begin to separate with the yellow carotene band getting ahead of the green chlorophyll band. When the extract has all been added to the column, add about 1/2 ml of hexane to the top of the column and drain to the sand. Then switch to the 90/10 mixture of hexane and acetone, adding enough of this solvent to completely remove the yellow band from the column. Continue collecting solvent in your waste solvent beaker until the yellow band has reached the bottom of the column and the solvent draining out begins to turn yellow. This yellow band is sometimes very narrow--don't miss it! When the drops are yellow replace the waste beaker with vial #1 and continue collecting in this vial until the drops lose their color or the vial is full. If the yellow band is not finished when the vial is full, continue to collect the yellow band in vial #2, 3, etc. When the yellow band is out of the column, collect any clear solvent in your waste beaker.

When the yellow band is out and the solvent you have most recently added has drained to the top of the sand, add your 70/30 mixture of hexane and acetone to move the green band further down the column. Continue to add this solvent mixture until all 10 ml have been added. Whenever the green color reaches the bottom of the column, remove your waste beaker and put in your next vial and subsequent vials as needed as you continue collecting the green band. When your green band first begins to collect, it may be quite pale. When all 10 ml of the 70/30 mixture have been added, you may need to add the straight acetone to further increase the polarity of the eluting solvent and bring out more of the chlorophyll band. When the major portion of the green band has been collected, replace the latest vial with your waste solvent beaker and allow the residual solvent to drain out of the pipette. This is another possible stopping point in the experiment.

<>Before thin layer chromatography (TLC), concentrate your colored solutions so that they will show up better on the TLC plates. A gentle stream of air or nitrogen in a hood is used to evaporate solvent so that only about 1/4 ml of yellow and 1/4 ml of green solution are left. If necessary, combine the contents of your vials as you evaporate so that you end up with just one vial of yellow carotenes and one vial of green chlorophylls. Also evaporate the original extract you saved for TLC so that no more than 1/4 ml is left. If a vial completely dries out, just add about 1/4 ml of hexane to reconsititute the solution. Your instructor will have a container available to discard the Pasteur pipette that was used as a column. Waste solvent should not be poured down the drain; place it in your laboratory's waste solvent container. Pipettes that were used to transfer solvents do not need to be washed and may be saved for later use. Green-stained pipettes should be discarded in the broken glass container.

Thin Layer Chromatography:

A thin layer chromatography (10 cm x 4 cm) plate is obtained and a horizontal pencil line is gently drawn about 1.5 cm from the bottom of the plate. On this line are spotted your three solutions (E, yellow and green) at equidistant intervals, using a separate microcap for each spot. Fill each capillary by dipping it in the solution and gently touch it to the plate to empty it. Use several short touches to empty each capillary so that the spots will be small. If the spots look light in color, re-spot on top of the original spots until the spots are fairly dark. Allow the spots to dry. Obtain a chromatography jar, place a filter paper upright against one wall of the jar and add about 10 ml of developing solvent (70% hexane-30% acetone). A little less than a centimeter of the filter paper should be standing in the solvent. Adjust the amount of solvent in the jar if necessary. Cap the jar and allow it to stand a few minutes until the filter paper becomes wet with solvent. Then place the TLC plate in the developing jar with the spotted end at the bottom of the jar, leaning the top of the plate against the filter paper. The bottom of the plate should be sitting in the solvent but the solvent should not be deep enough to submerge the spots. The sides of the TLC plate should not touch the filter paper. Cap the jar and allow the solvent to rise up the plate undisturbed until about 80% of the plate is wet. Remove the plate and quickly draw a pencil line across the plate to mark the farthest reach of the solvent. This is called the solvent front. Allow the wet plate to dry in the hood. Visible spots should be circled in pencil since the colors may fade over time.

Retention fractions (R_f):

Different compounds should rise to different heights on your TLC plates, however the exact height a particular compound rises depends on how high the solvent is allowed to rise up the plate. If the solvent travels higher, then the spots all travel higher too. To correct for this difference and generate a number which can be compared to reported values or to other people’s work, the retention fraction or R_f value is calculated. The retention fraction is defined to be the fractional rise of the spot compared to the rise of the solvent. The R_f value for a compound will change if a different developing solvent or a different type of plate is used. Calculate R_fvalues for each spot on your plate. Spots with the same R_fvalues within experimental error and the same appearance should be the same compound.

If you had just two components in your original extract, this is what your results might look like:

<>A sketch of your plate should be included in your notebook. Your plate may be considerably more complex in appearance than the example. Be sure to clearly label the lanes as coming from extract, yellow band or green band and label the colors of as many of the separated spots as possible. Identify as many spots as you can. Determine which pigments were present in the yellow band and which were present in the green band from your column. In your conclusion you should address how well the column separated the various pigments. Depending on the leaf sample, the conditions of the experiment, and how much sample was spotted on the tlc plate, you may observe other pigments. These additional components can result from air oxidation, hydrolysis, or other chemical reactions involving the pigments discussed in this experiment. In the crude extract, you may be able to see the following components (in order of decreasing R_f values):

Carotenes (1 spot) (yellow-orange)

Pheophytin a (gray, may be nearly as intense as chlorophyll b)

Pheophytin b (gray, may not be visible)

Chlorophyll a (blue-green, more intense than chlorophyll b)

Chlorophyll b (green)

Xanthophylls (possibly 3 spots: yellow)

koster.sand@uwlax.edu

last modified 10/3/08