CHAPTER 8 – DIVERSITY OF PHOTOSYNTHETIC PIGMENTS



DIVERSITY OF PHOTOSYNTHETIC PIGMENTSPaul R. EcklundLABORATORY SYNOPSISThis laboratory continues your study of protists. In this laboratory you will extract photosynthetic pigments from organisms representing three phyla of algae and one plant phylum. Then you will use thin-layer chromatography to separate the pigments in each extraction mixture, and identify the characteristic pigments of each phylum. In your discussion/interpretation of the chromatograms, you will consider the principles of thin-layer chromatography and evolutionary relationships among the algae and plants. The background information provided for this laboratory deals not only with the diversity of pigments found in several groups of photosynthetic organisms, but also discusses the origin and evolution of those groups of organisms.LABORATORY OBJECTIVESConceptualAt the end of this laboratory and the associated laboratory lecture you should1.know the distinctive photosynthetic pigments of the brown, red and green algae.2.be able to recognize the algal specimens you used and know which phylum each specimen is in.3.know what endosymbiosis is, and the proposed origin and evolution of chloroplasts according to the endosymbiotic hypothesis for eukaryotic cell evolution.4.understand the basic concepts and principles of thin-layer chromatography (for example, molecular polarity, adsorption, adsorbent, stationary and mobile phases, solvent system).5.know which atoms and atom groups cause polarity in photosynthetic pigment molecules.6.understand the general procedures of thin-layer chromatography.7.understand how pigment analysis can be used in the classification of photosynthetic organisms.8.appreciate the toughness of seaweeds.9.appreciate the “grind” of extracting pigments from algae by hand power.10.understand what is meant by the red and green lineages in chloroplast evolution.ProceduralAt the end of this laboratory you should1.learn a grinding technique for extracting substances from algae and plants.2.observe/learn a partitioning procedure for concentrating a solution.READING ASSIGNMENTTo prepare for this laboratory read:In Castro and Huber, Marine Biology, 9th ed. (2013):Chapter 5:pages 86-89 (Prokaryotes to Archaea)pages 93-97 (Unicellular Algae)Chapter 6:pages 102-114 (Red algae and green algae . . . to end of chapter)In Diversity of Photosynthetic Pigments:pages INTRODUCTION, OVERVIEW OF ACTIVITIES, THIN-LAYER CHROMATOGRAPHY sections.QUESTIONS TO PREPARE YOU FOR THIS LABORATORY1.What important reproductive characteristic distinguishes multicellular algae from plants?2.Which pigments are found in all organisms that perform oxygen-producing photosynthesis?3.According to the endosymbiotic hypothesis for the origin and evolution of eukaryotic cells, what was the precursor of the chloroplast?4.In the thin-layer chromatography system you will use, what is the stationary phase?5.a.What is adsorption?b.What property must a pigment molecule have to adsorb to the stationary phase of a chromatography system?6.In thin-layer chromatography, what determines how fast a certain kind of molecule will move relative to the movement rate of the mobile phase?INTRODUCTIONAll algae, all plants and two phyla of bacteria use chlorophyll a and other pigments to perform oxygen-evolving photosynthesis. The term “algae” applies to all eukaryotic, photosynthetic organisms that are not plants. Certainly, the distinction between plants, which are multicellular, and the unicellular algae is obvious. From your assigned reading, however, you should appreciate that some types of multicellular algae, particularly in the red and brown algae, may be larger than many plants and structurally as complex as some simpler plants. The major distinguishing difference between complex, multicellular algae and plants is in their treatment of a zygote formed by the union of two gametes in sexual reproduction. Algae release zygotes shortly after they are formed, and thus provide no protection nor nourishment for the organism that develops from each zygote. Plants retain their zygotes in multicellular female reproductive structures, which give protection and nutrients to the developing embryos.One criterion used to determine evolutionary relationships among the algae and classify them is the pigment composition of their chloroplasts. Figure 8.1 shows the chemical structures of several chloroplast pigments.Molecules of chlorophyll a attached to specific proteins function as the catalytic pigment in the two photosystems of all algae and plants. Most of the chlorophyll a in the photosystems, however, is used to capture photons and/or transfer energy to the catalytic forms of the pigment. All pigments other than chlorophyll a, which assist in absorbing photons to energize the photosystems, are regarded as accessory pigments. Chlorophyll b and the two forms of chlorophyll c are green accessory pigments. These chlorophyll types are not common to all the algae.All algae and plants possess one or more kinds of carotenoid pigments (Figure 8.1 D, E and F). Carotenoids collectively include the hydrocarbon carotenes (for example, beta carotene, Figure 8.1 D) and their hydroxyl derivatives, the xanthophylls. These pigments are yellow, orange and red and absorb light in the blue-green part of the spectrum. Some of the carotenoids assist in harvesting light for the photosystems. For example, the xanthophyll fucoxanthin (Figure 8.1 F) is the main accessory pigment in diatoms, the golden algae and the brown algae; its relative abundance in their chloroplasts gives these algae their characteristic color. A major role of some of the carotenoids is protection of the photosystems by absorbing blue and near ultra violet radiation that can damage chlorophyll molecules and other components.The endosymbiotic hypothesis is a currently popular and well-supported hypothesis which explains the origin of chloroplasts in eukaryotic cells. This hypothesis proposes that prokaryotic, oxygen producing, photosynthetic cells were “eaten” by phagocytic cells. Instead of being digested, however, the ingested cells lived as endosymbionts within the host cells, coordinated their reproduction with that of the host cells, and eventually evolved into chloroplasts. In the evolution of the endosymbiont to a chloroplast, much of the endosymbiont’s genetic material was transferred to the host cell’s nucleus, making the endosymbiont dependent on the host cell.Ancient members of the prokaryotic phylum Cyanobacteria (blue-green bacteria) are assumed to have been the endosymbionts that evolved into chloroplasts in the first algae. This assumption is based on numerous structural, biochemical and genetic similarities between cyanobacteria and the chloroplasts of photosynthetic eukaryotes. Furthermore the fossil record indicates that cyanobacteria were well established on earth when the first photosynthetic eukaryotic cells originated.Ancient endosymbiotic cyanobacteria cells apparently evolved with relatively few changes into the chloroplasts of the eukaryotic phylum Rhodophyta (red algae) because the cells of blue-green bacteria and the chloroplasts of red algae are very similar in structure and pigment composition. Biliproteins are the dominant accessory pigments in members of these two phyla. The blue-green phycocyanins and the red phycoerythrins consist respectively of the pigment molecules phycocyanobilin and phycoerythrobilin covalently bonded to specific proteins (see Figure 8.1C). Only the members of these two phyla possess various forms of phycocyanin and phycoerythrin arranged into complex light harvesting structures called phycobilisomes which are attached to the surface of the thylakoids.The green algae (phylum Chlorophyta) have been on earth at least as long as the red algae. The chloroplasts of green algae differ from the blue-green bacteria by possessing chlorophyll b and lacking biliproteins as accessory pigments. If ancient endosymbiotic blue-green bacteria evolved into the chloroplasts of green algae, they lost the biliproteins and substituted chlorophyll b in the process. In the late 1960s a photosynthetic, oxygen-producing green bacterial species possessing chlorophylls a and b and lacking biliproteins was identified. Additional species of this bacterial type have since been identified. These green bacteria have been assigned to the phylum Prochlorophyta or Chloroxybacteria (Margulis and Schwartz 1988). Members of the genus Prochloron are unicellular and share many characteristics with the chloroplasts of green algae. The discovery of these prokaryotic “counterparts” to the chloroplasts of green algae suggested that an ancient green, oxygen producing bacterial type might have been the precursor of the green chloroplast lineage. The results of recent research, however, more strongly support a cyanobacterium (not a green bacterium) precursor of green chloroplasts.Chlorophyll b is the major photosynthetic accessory pigment in all the green algae, the photosynthetic euglenids and all plants. When systematists observe a trait that is shared by different groups of organisms, they hypothesize that the trait evolved only once and it is shared because the groups inherited it from a common ancestor; that is, the groups are related by their evolutionary histories. Groups of organisms which share many features are assumed to be more closely related than groups with fewer common features. The Chlorophyta and plants have several common features including chloroplasts with a double membrane envelope, starch as the major food reserve and cellulose as the principal cell wall constituent. Furthermore, recent studies indicate the presence of many similarities in their genetic information. Collectively, these shared features provide strong evidence for a close relationship between the Chlorophyta and plants. It is assumed that certain ancient species of green algae gave rise to plants. The evidence for a common ancestor of green algae and plants is now so strong that many systematists wish to include the green algae and plants in a clade. The tentative clade is named Viridiplantae in Campbell and Reece (2005).In diatoms, golden algae and brown algae chlorophyll c, instead of chlorophyll b, is the accessory chlorophyll and fucoxanthin is the predominant carotenoid in the chloroplasts. Furthermore, the chloroplasts are unlike those in other phyla by having two additional membranes around the double membrane envelope; the outer membrane is continuous with the outer membrane of the nuclear envelope. The major food reserve in these algae is a polysaccharide called laminarin. These shared traits imply an evolutionary relationship among these algal groups; presumably they have a common ancestor. The complexity of the chloroplast with its adjacent membranes has caused some evolutionary biologists to propose that this type of chloroplast originated from a eukaryotic endosymbiont.Table 8.1 summarizes the distribution of chloroplast pigments in the various groups mentioned above.Table 8.1. Pigments in the thylakoid membranes of various groups which perform oxygen-evolving photosynthesis. An X indicates the presence of the pigment(s). Phyla whose names are in bold type are represented in this laboratory.Figure 8.1 (above and next page). Structures of various chloroplast pigments. Lines represent covalent bonds. In the ring diagrams, carbon atoms are at the line junctions without atom symbols. A common feature of the chlorophylls and the phycobilins (A-C) is a structure called a tetrapyrrole which consists of four rings, each composed of four carbons and a nitrogen, linked together. The tetrapyrrole is linear in the phycobilins, but in the chlorophylls it is a ring with a Mg2+ ion in the center. Chlorophyll c does not have the long hydrocarbon chain possessed by chlorophylls a and b. In a photosynthetic system, the chlorophylls and phycobilins are conjugated with proteins. D, E and F are carotenoids; E and F are xanthophylls. A common feature of all these pigments is a series of alternating single and double bonds between the atoms. This bonding arrangement enables these molecules to absorb photons and remain chemically stable. Diagrams were adapted from Salisbury and Ross (1969) and South and Whittick (1987).Figure 8.1 continued.OVERVIEW OF LABORATORY ACTIVITIESIn this laboratory you and your labmates will extract in acetone the chlorophylls and carotenoids from a selected species of each of the algae phyla Rhodophyta, Phaeophyta and Chlorophyta and the plant phylum Magnoliophyta (flowering plants) (Anthophyta in Table 8.1). You will use a technique called thin-layer chromatography (explained below) to separate and identify the acetone-extracted pigments from each species. Then you will compare the developed chromatograms and look for similarities and differences in the chloroplast pigment composition of the different species.THIN-LAYER CHROMATOGRAPHYChromatography, in its various forms, is used to separate different molecular species from a mixture. The name "chromatography," which means "color writing," was given first to a technique used to separate chloroplast pigments. Now the term applies to numerous techniques used to separate different kinds of molecules, whether they are colored or not. For example, Melvin Calvin and associates employed a form of chromatography to separate and identify the colorless intermediates of the "Calvin cycle" of photosynthesis after they were extracted collectively from a green alga.All chromatographic systems consist of a stationary phase and a mobile phase (a liquid or gas which moves through the stationary phase). Molecular species are separated by their relative affinities for the two phases. Molecules with a strong affinity for the mobile phase and little attraction for the stationary phase readily move with the mobile phase and experience little hindrance to movement by the stationary phase. On the other hand, molecules that have a weaker affinity for the mobile phase and are strongly attracted to the stationary phase bind reversibly to the latter, inhibiting their rate of movement in the mobile phase.Thin-layer chromatography (TLC) is often regarded as a microchromatographic technique because the period of time required for separation of different molecular species is relatively short, and relatively small amounts of substances can be separated and detected. TLC entails a process called adsorption which is the adhesion of molecules to the surface of a substance called an adsorbent. Adsorption results from the electrostatic attraction between the mobile molecules and the adsorbent material. The stationary phase of our TLC system is the adsorbent, a thin-layer (100 ?m in depth) of silica gel (specially prepared silicic acid) bonded to a thin sheet of inert plastic material. The mobile phase is the solvent system consisting of a mixture of toluene and acetone in a volume ratio of 3:2, respectively.In your use of TLC, you apply a small amount of a solution of extracted pigments as a spot near one end of an adsorbent sheet. Then you put the "spotted" sheet vertically into a test tube containing a small volume of the solvent system and stopper the tube. The solvent system moves up the thin-layer of adsorbent by capillary action. As the solvent system moves up the stationary phase, it picks up the applied pigments and transports them up the adsorbent sheet. The rate at which each chloroplast pigment molecule migrates in the mobile phase depends upon its relative attraction for the stationary phase and for the two different molecular species in the mobile phase.Figure 8.2. Chemical structures of toluene and acetone (mobile phase).The stationary phase is a lattice of silica molecules, which are highly polarized. Polar regions of pigment molecules are electrostatically attracted to the silica. The mobile phase consists of toluene, which is nonpolar, and acetone, which is polar (Figure 8.2). Toluene attracts nonpolar regions of pigment molecules and acetone has an affinity for both polar and nonpolar parts of the molecules. Since all of these attractive interactions are reversible, the pigment molecules are continually attaching to the adsorbent and being pulled off by the migrating solvent components. The following are empirical generalizations regarding the binding of molecules to the silica gel (stationary phase).H H | |1.Saturated hydrocarbons (e.g., H–C–C...) are not bound, or are only weakly bound. | |H H2.The binding of unsaturated hydrocarbons increases with the number of double bonds and the number that are in conjugation (alternating single and double C-C bonds; see Figure 8.1D-F).3. Molecular groups substituted for hydrogens in a hydrocarbon increase the binding affinity of the molecule.The relative effect of various groups on binding affinity are given below.–O–alkyl* < –C=O < –NH2 < –OH < –COOH*An alkyl group is a hydrocarbon consisting of one or more carbon atoms saturated with hydrogen atoms.MORPHOLOGY OF THE SELECTED ALGAL SPECIESObserve the external morphology of the specimens representing three algae phyla. All three of the specimens are marine algae from the coast of Maine.The body of a multicellular alga is called a thallus. A complete thallus of any of the three specimens displayed has a basal structure, called a holdfast, to attach it to a firm surface.Note that each specimen is very flexible, but also very tough with a high resistance to tearing. (You will appreciate the toughness of these thalli when you attempt to grind them.)?How are these traits adaptive for these organisms in their natural environment?PHYLUM RHODOPHYTA (Red Algae): Porphyra sp.The thallus is a thin sheet composed of only two cell layers. The specimen may appear more brown than red. The color of the thallus depends on the ratio of its pigments. Porphyra exhibits a wide range in color depending on the light intensity of its environment. Organisms growing in shallow water with high light intensity may be dull green. As the water depth of its environment increases (and light intensity decreases) Porphyra's phycoerythrin content increases, and it becomes more reddish.Porphyra is used as a human food. The Japanese have farmed seabeds of this red alga for many years. Dried sheets of Porphyra thalli are called nori, which is used in various soups and sauces and for the wrapping of sushi. PHYLUM CHLOROPHYTA (Green Algae): Ulva lactucaThe thallus is very similar to that of Porphyra–a thin sheet of only two cell layers. The common name for Ulva is "sea lettuce."PHYLUM HETEROKONTOPHYTA (Brown Algae): Ascophyllum nodosumThe thallus is highly branched. Each branch has at least one oblong bladder filled with gas. You may have seen this alga in the seafood section of a market. Fresh seafood is often packed for shipping in Ascophyllum nodosum and displayed on it in the store. Your specimen may have reproductive structures, called receptacles, along each branch. Each receptacle is a yellow-green, oblong, pea-sized structure attached by a short stalk. You also may see small, dark red or brownish tufts of the red alga Polysiphonia lanosa growing on some of the ANIZATION OF LABORATORY ACTIVITIESYou will work in pairs (or groups of 3) to do the extraction and concentration procedures. Each pair/group of students extracts chloroplast pigments from one alga or spinach. You will work in groups of 3 to do the chromatography procedures. You will be assigned one of the following extractionsRed alga (Porphyra): group of 4 studentsGreen alga (Ulva): group of 3 studentsBrown alga (Ascophyllum): group of 4 studentsAngiosperm (Spinacea): group of 3 studentsEach group obtain the required materials for your extraction procedure. See the next section.NOTE:Containers of acetone and petroleum ether are distributed on the laboratory benches. Laboratory sand and anhydrous sodium sulfate are on the bench closest to the blackboard on the west end of the Zoology lab.REQUIRED MATERIALS FOR EXTRACTION PROCEDURES1 mortar and pestle1 10 ml graduated cylinder1 metal test tube rack1 small test tube (which must be thoroughly dry inside)1 stirring rod2 50 ml glass beakers (thoroughly dry inside; dry with a paper towel, if necessary)1 50 ml or 100 ml glass beaker (thoroughly dry inside; dry with a paper towel if necessary)1 5 ml pipette1 pipetting aid1 microcentrifuge tube (small, plastic, tapered test tube)1 microcentrifuge tube rack1 marking pen1 package of algal material or spinach leavesPIGMENT EXTRACTION PROCEDURESUlva, Porphyra, or Spinach1.Remove all of the algal “sheets” or spinach leaves from the plastic bag and spread them on a paper towel.2.Blot "sheets" of the alga (or spinach leaves) with additional paper towels until all surface water is removed (excess water interferes with the extraction); then tear the tissue into pieces about 1 x 1 cm and place them in the mortar.3.Add about 1/2 teaspoon of sand and 8 ml (use graduated cylinder) of acetone to the mortar.4.Hold the mortar firmly and with the pestle grind the tissue vigorously until it is thoroughly macerated and the acetone is dark green (mash down on the tissue and then grind it against the side and bottom of the mortar). Consult your lab instructor regarding the thoroughness of your extraction before you proceed to the next step.5.Carefully pour off the solution into the glass beaker. (If there is too little extract to pour, add 2-3 ml of acetone and then pour.)6.Add an additional 4 ml of acetone to the mortar and grind the tissue again until the extract is dark green. Then pour the extract into the same beaker. You may need to use the pestle to press out as much extract as you can from the ground tissue.7.Go to PROCEDURES FOR CONCENTRATING THE PIGMENT EXTRACT FOR CHROMATOGRAPHY.Ascophyllum1.Each member of the group obtains a razor blade and a small cutting board.2.Remove all of the algal material from the plastic bag and spread it on a paper towel. If receptacles are present on the thallus, cut them off and discard them in the trash. Gelatinous material in the receptacle interferes with the extraction.3.Blot the algal material with paper towels until all surface water is removed; excess water interferes with the extraction.4.Each person with a razor blade and cutting board take a portion of the algal clump, arrange the branches to be side-by side on the cutting board and chop the branches into approximately 3 mm (1/8 in) sections. Chop all the material provided and dump the pieces into the mortar.5.Add about 1 teaspoon of sand and 8 ml (use graduated cylinder) of acetone to the mortar, hold the mortar firmly and grind vigorously with the pestle until the acetone is dark green. (Mash down on the tissue, then grind it against the side and bottom of the mortar.) Consult your lab instructor regarding the thoroughness of your extraction before you proceed to the next step.6.Carefully pour off the liquid into the glass beaker. (If there is too little extract to pour, add 2-3 ml of acetone and then pour.)7.Add an additional 4 ml of acetone to the mortar and grind the tissue again until the acetone is dark green. Pour the extract into the same beaker. You may need to use the pestle to press out acetone from the ground tissue.8.Go to the PROCEDURES FOR CONCENTRATING THE PIGMENT EXTRACT FOR CHROMATOGRAPHY.PROCEDURES FOR CONCENTRATING THE PIGMENT EXTRRACT FOR CHROMATOGRAPHY1.Allow the contents of the glass beaker to settle.2.Carefully decant the acetone portion of the beaker’s contents in to a clean, dry 50 ml beaker. Be sure to decant only the clear acetone extract which forms the upper layer of the fluid in the beaker. Do not pour out the sediment or the cloudy solution in the bottom layer.3.Use the graduated cylinder to add 3 ml of petroleum ether to the acetone extract in the beaker.4.Use the stirring rod to thoroughly mix the petroleum ether and the acetone extract, then carefully pour the mixture into a dry, small test tube.Observe the separation of the liquid phases. The upper layer is a solution of pigments in petroleum ether with some acetone; the bottom layer is acetone with some water, and may contain some pigments.NOTE: If you are NOT preparing an extract from Porphyra, progress to step 6. If you are preparing an extract from nori, do the special procedure below before going to step 6.Special Procedure for Extract from Porphyra (Nori)The extract from Porphyra yields a green upper layer consisting of chlorophylls and carotenoids in a mixture of petroleum ether and acetone and a lower layer, which may be pink, red, purple or blue, consisting of phycoerythrin/phycocyanin in a mixture of acetone and water.Allow your lab instructor and labmates to see these two colored layers before you do step 6.Continue with steps 6-11, but do not discard the colored acetone-water layer in the test tube (step 12). Give the test tube containing the acetone-water layer to your lab instructor.6.Allow the layers to separate for 3-4 minutes, then use a 5 ml pipette with a pipetting aid to remove very carefully most of the petroleum ether from the test tube (leave the bottom 2 mm and do not remove any of the lower layer). Quickly dispense the petroleum ether into a clean and dry 50 ml beaker.7.Add another 3 ml of petroleum ether (use a graduated cylinder) to the contents in the test tube and thoroughly mix with a stirring rod. Repeat step 6 using the same pipette. Add the removed petroleum ether to that in the beaker used in step 6.8.Use the microspatula to add one level scoop of anhydrous sodium sulfate to the petroleum ether in the beaker. Recap the sodium sulfate container immediately after you use it.NOTE: The anhydrous sodium sulfate will not dissolve; it removes water from the solution.9.Under the hood gently swirl the beaker until the volume of petroleum ether has been reduced to about 0.5 ml (just enough to cover the bottom of the beaker). If you inadvertently allow all the petroleum ether to evaporate, add 0.5 ml of petroleum ether to the beaker and gently swirl it to dissolve the pigments.10.Carefully pour the remaining pigment solution into a microcentrifuge tube and label the tube with a letter indicating the source of the extracted pigments. B = brown algaG = green algaR = red algaS = spinach11.Put the microcentrifuge tube in the tube rack, and put a capillary tube in the microcentrifuge tube.12.Discard the acetone in the Hazardous Waste bottle in the hood. Discard the ground tissue and sand in the mortar into the special waste container in the hood. Please do not pour acetone down the drain.CHROMATOGRAPHY PROCEDURESWork in groups of 3 or 4.Each group member “spots” a chromatography strip with a pigment extract from one of the 4 different sources, and develops the chromatogram. Each group should have chromatograms with pigments from the 4 different sources (3 algae and 1 plant), so groups must share extracts.Each group member selects the source of pigments he/she will use (brown, green or red alga or spinach).Each group obtains the following items:4 large test tubes with stoppers (these tubes must be completely dry inside)1 wooden test tube rack for the large tubes1 clean 1 ml pipette1 pipetting aid2 forceps2 rulersPlace the test tubes in a test tube rack. Use the 1 ml pipette with a pipetting aid to dispense 0.8 ml of chromatography solvent (we may need to add more solvent!) into each tube. Stopper each tube immediately after the solvent is added, and keep the tube stoppered until a chromatography strip is put in it.Procedure for Preparing Chromatography StripsPrecaution: The adsorbent surface of the TLC strip must not be scratched, and it must not be touched directly by fingers; skin oils interfere with the movement of the solvent system through the adsorbent. If the adsorbent becomes wet with water, it is ruined!1.Place a clean, flat paper towel on your work surface. Your instructor will use forceps to lay four strips, dull side up/shiny side down, on the paper towel.2.Use a pencil (not a pen) to make 2 faint marks on one long edge of the strip–one at 0.5 cm from the bottom and the other at 0.5 cm from the top edge. (If you must hold down the strip to do this, put a piece of paper between your fingers and the adsorbent.) Near the top edge, write the letter indicating your pigment source (B, G, R, or S).3.Obtain the appropriate pigment extract. Allow the capillary tube in it to fill by capillary action, then place your finger firmly over the upper end of the tube and keep your finger there while you apply extract to the TLC strip to control the flow of the liquid.Precaution: If a droplet of extract forms on the tip of the capillary tube, gently blot it on a paper towel before you apply extract to the chromatography strip.4.Apply a very small spot of extract in the middle of the width of the strip at 0.5 cm from the bottom. Gently touch the lower tip of the capillary tube to the strip so that only a small amount of extract is wicked out. Allow the spot to dry completely (about 10 sec), then apply another spot on top of the first one. Repeat this application procedure 5 times. (You must open the top of the capillary tube to fill it, but remember to keep it closed while applying extract to the strip.)5.After completing your application of extract to the TLC strip, allow the spot to dry completely before placing the strip in the chromatography tube. If the spot is dry, it will be dull; if it is still wet it will be shiny. You can hasten the drying of the spot by giving it a gentle stream of air from the air jet on the bench top.Developing the ChromatogramsPrecaution: Leave the chromatography tube in the test tube rack when you put the TLC strip in it and during the development of the chromatogram. Movement of the chromatography tube may cause the solvent system to move unevenly up the adsorbent sheet.1.Use forceps to carefully hold the top of the TLC strip and place the strip into the chromatography tube. Then gently stopper the tube.2.Observe the movement and separation of pigments as the solvent front moves up the TLC strips. Remove the strip from the tube when the solvent front is 0.5 cm from the top of the strip. Place the chromatogram on a paper towel to dry.3.Discard the TLC solvent in the special waste container in the hood. Do not wash or rinse the chromatography tubes; place them upside down on the pegs of the rack in the hood. Put the stoppers in the labeled container.CLEAN UP PROCEDURES1.Pour the solvent from the chromatography tubes into the Hazardous Waste bottle in the hood. Do not wash the chromatography tubes; place them upside down on the pegs of the rack in the hood. Put the stoppers in the labeled container.2.Pour the pigment extract (if any remains) from the microcentrifuge tubes into the Hazardous Waste bottle, then put the microcentrifuge tubes in a designated container also in the hood.3.Put capillary tubes in the “broken glass” containers by the sinks.4.Put pipettes, tips up, in the “used pipettes” container.5.Wash mortars, pestles and all remaining glassware and plasticware in soapy water; rinse them in tap water and then distilled water and place them inverted on drain racks/hang them on drain pins to dry. Use a scrubbing pad and soapy water to thoroughly remove pigment stains from the grinding surfaces of mortars and pestles.6.Return all the other equipment to the side bench.7. Discard chromatograms in the trashcans.REFERENCES AND SUGGESTED READINGCampbell NA, Reece JB. 2005. Biology. 7th ed. San Francisco, CA: Pearson-Benjamin/Cummings.deDuve C. 1996 (April). The birth of complex cells. Scientific American 274(4):50-57.Falkowski PG, and others. 2004. The evolution of modern eukaryotic phytoplankton. Science 305:354-360.Lee TF. 1977. The seaweed handbook. Boston, MA: The Mariners Press.Margulis L, Schwartz KV. 1988. Five kingdoms. 2nd edition. New York: W.H. Freeman and Co.Motten AF. 1995. Diversity of photosynthetic pigments. In: Goldman CA., editor. Tested studies for laboratory teaching: proceedings of the 16th workshop/conference of the Association for Biology Laboratory Education (ABLE); 1994 June 7-11; Emory University, Atlanta, GA. Toronto, Ontario: Association for Biology Laboratory Education p. 81-98.Randerath K. 1966. Thin-layer chromatography. 2nd edition. New York: Academic Press. Translated from German by D.D. Libman.Salisbury FB, Ross CR. 1969. Plant physiology. Belmont, CA: Wadsworth Publishing Co., Inc.South GR, Whittick A. 1987. Introduction to phycology. Oxford: Blackwell Scientific Publications. ................
................

In order to avoid copyright disputes, this page is only a partial summary.

Google Online Preview   Download