Photosynthesis


Plants Do Photosynthesis

Photosynthesis is the process of converting light energy to chemical energy and storing it in the bonds of sugar. This process occurs in plants, some algae (Kingdom Protista), and the cyanobacteria (also known as “bluegreen algae,” Kingdom Monera). Photosynthesis requires only light energy, CO2, and H2O to make sugar. The process of photosynthesis takes place in the chloroplasts, specifically using chlorophyll, the green pigment involved in photosynthesis.

Leaf Cross-Section Photosynthesis takes place primarily in plants’ leaves, and little to none occurs in stems, etc. The parts of a typical leaf include the upper and lower epidermis, the mesophyll, the vascular bundle(s) (veins), and the stomates. The upper and lower epidermal cells do not have chloroplasts, thus photosynthesis does not occur there. They serve primarily as protection for the rest of the leaf. Chloroplast The stomates are holes which occur primarily in the lower epidermis and are for air exchange: they let CO2 in and O2 out. The vascular bundles or veins in a leaf are part of the plant’s transportation system, moving water and nutrients around the plant as needed. The mesophyll cells have chloroplasts and this is where photosynthesis occurs.

As you hopefully recall, the parts of a chloroplast include the outer and inner membranes, intermembrane space, stroma, and thylakoids stacked in grana. The chlorophyll is built into the membranes of the thylakoids.

The light reaction happens in the thylakoid membrane and converts light energy to chemical energy. This chemical reaction must, therefore, take place in the light. Chlorophyll and several other pigments such as beta-carotene are organized in clusters in the thylakoid membrane and are involved in the light reaction. Each of these differently-colored pigments can absorb a slightly different color of light and pass its energy to the central chlorphyll molecule to do photosynthesis.


Pigments Involved in Photosynthesis

In this lab you will be examining the pigments present in plant leaves, separating/isolating these pigments from each other, and determining absorption spectra for each of them.

Chlorophyll A (chloro = green, phyll = leaf) is the pigment used by plants to convert energy from the sun into chemical energy useful to the plant, but other pigments present in leaves also help to “harvest” light energy. This energy is stored by converting carbon dioxide and water to sugar. The chemical reaction for this is 6 CO2 + 12 H2O (+ light energy) → C6H12O6 + 6 O2 + 6 H2O. Central Structure of Chlorophyll This sugar is stored by the plant as starch (thus the occurrence of photosynthesis could be demonstrated using the iodine test for starch). Benedict’s solution could be used to test for the presence of sugar (usually found in the leaf veins, indicating transfer of sugar from one part of the plant to another).

The central part of the chemical structure of a chlorophyll molecule is a porphyrin ring, which consists of several fused rings of carbon and nitrogen with a magnesium ion in the center.

Chlorophyll looks green because it absorbs red and blue light, making these colors unavailable to be seen by our eyes. It is the green light which is NOT absorbed that finally reaches our eyes, making chlorophyll appear green. However, it is the energy from the red and blue light that is absorbed that is, thereby, able to be used to do photosynthesis. The green light we can see is not/cannot be absorbed by the plant, and thus cannot be used to do photosynthesis.

Beta-Carotene and Vitamin A
Structure of Vitamin A and β-Carotene
Besides chlorophylls A and B, various other pigments, including carotenes (carot = carrot), xanthophylls (xantho = yellow), and anthocyanins (antho = a flower, cyano = blue, dark blue), are often found in plant leaves. The chemical structures of these molecules are illustrated in many organic chemistry and cell physiology books. Because of their different colors, many of the carotenes and xanthophylls are capable of “capturing” solar energy that the chlorophyll cannot and transferring that energy to the chlorophyll enabling photosynthesis to occur. Anthocyanins are not involved in photosynthesis.


Paper Chromatography

Paper Before and After Once a mixture of these pigments has been extracted from a leaf together, because each of these pigments, including the chlorophylls, has a different chemical structure and formula, the mixed pigments can be separated from each other be a process known as paper chromatography (chromo = color; graph = to write). In this process, the mixed pigments are dissolved in a mixture of two (or more) solvents and allowed to soak into a piece of paper by capillary action. Typically, one of the solvents used is more covalent while the other is more polar or ionic, and their molecular weights differ considerably. Various of the leaf pigments, thus, are more, or less, soluble in the different solvents, so as the solvent system wets the paper, the various pigments move into/across the paper at various rates depending on their sizes (molecular weight), relative number of covalent or ionic bonds in the molecule, and other factors based on their chemical structures: normally, the smallest move fastest and farthest.

Once the pigments are separated, a tentative identification of each may be made (to be confirmed by obtaining an absorption spectrum of each.). Chlorophyll A appears as a blue-green band while chlorophyll B is a yellow-green band. Carotenes are bright yellow to orange while xanthophylls are a slightly greenish yellow. Anthocyanins are reddish, violet, or blue, and are not soluble in organic solvents, thus typically do not move up the chromatogram at all.


Light Absorbed by Each of the Pigments

The fact that each of these pigments appears as a different color is an indication that each is absorbing different wavelengths of light. Remember the color(s) that we see is whatever the plant has NOT absorbed (For example, chlorophyll A looks green because it is not absorbing and using green light). According to the literature, chlorophyll A has two absorption peaks (absorbs the most light) at around 428 nm (blue-violet range) and at around 660 to 700 nm (red range), while chlorophyll B absorbs best at around 453 and 643 (to 650) nm. Beta-carotene, the most common carotene (and precursor of vitamin A), has an absorption peak at a wavelength of 451 nm (at the blue-violet end of the spectrum). Each of these pigments or the mixture as extracted from the leaf can be examined with a spectrophotometer to determine its absorption spectrum, thus confirming its identity.


Effects of Wavelength on Plant Growth

While the present experiment will not test this, it has been reported that the blue/violet light absorbed by plants is responsible for foliage growth. Plants grown in only blue light are compact with lush, dark-green leaves but few flowers. Red and far-red light affect the growth processes (elongation and expansion) in various plant parts. Consequently, these colors are responsible for flower development among other things. Incandescent light is a good source of light in the red range but lacks output in the blue range while fluorescent light tends to be better in blue and lacking in the red end of the spectrum. Vitalightr and similar special fluorescent-type bulbs are specially-designed for high output in both the blue and red ranges, yet are low in the green range (hence tend to look purplish or pinkish in color). Many plant growers combine incandescent and fluorescent lights.


Safety Considerations

The chemicals in the solvent system being used in Part A and the ethanol used in Part B are flammable, thus should be kept away from any open flames. Pouring of solvent system to/from the Erlenmeyer flask should take place in a fume hood, and the reagent bottle and your flask should immediately be capped. It probably is also a good idea to not breathe too much of it.


Materials Needed

Part A — Paper Chromatography

Part B — Spectra of Pigments


Procedure

Part A — Paper Chromatography

    solvent in flask
    solvent in flask
  1. A 250-mL Erlenmeyer flask, #8 stopper, and T-pin should be obtained. Working in the fume hood, a few millimeters (depth) of the 90% petroleum ether + 10% acetone solution should be poured into the bottom of the flask and the stopper placed on the flask so that the “fumes” can start to accumulate in the flask while the next few steps are performed. The air in the flask must become saturated with fumes from the solvent or the chromatography won’t run properly.
  2. ready to measure paper
    ready to measure paper
    make paper this long
    make paper this long
    fold paper here
    fold paper here

    second fold here
    second fold here
  3. A piece of chromatography paper slightly longer than what will fit into the flask should be obtained. It is important to touch it as little as possible, preferably only at the edges. The bottom edge of the paper should be cut as straight as possible. The paper should be long enough that when a T-pin is used to attach it to the underside of the stopper, the tip of the paper will reach to within a few millimeters of the bottom of the Erlenmeyer flask. This paper must be kept as clean as possible, handled only by the edges, and only set on clean surfaces. If it is necessary to mark on the chromatography paper, only pencil should be used, not pen (ballpoint ink is alcohol-soluble).
  4. spinach, penny, and paper
    spinach, penny, and paper
    line on paper
    roll line on paper

    darker line on paper
    darker line on paper
  5. A spinach (or other) leaf should be obtained. The chromatography paper should be laid on a piece of clean paper and the leaf laid over the chromatography paper. The edge of a penny (or other coin) may be used to roll (smash) a stripe of color across the paper about 1.5 to 2 cm above the end. The leaf should be moved so a new portion of the leaf is over the stripe, and re-rolled with the penny over/onto the same place on the paper to darken the stripe. This process should be repeated several times as needed to obtain a dark stripe. The stripe should be allowed to dry before proceeding.
  6. pin here, view 1
    pin onto stopper, view 1
    pin here, view 2
    pin onto stopper, view 2
  7. The chromatography paper should be held up next to the flask to judge the exact length of paper needed such that when the paper is pinned to the bottom side of the stopper, the bottom end of the paper will be just below the surface of the solution. If needed, the top end of the strip should be folded over at the right place. The flask should not be left open while pinning the paper to the stopper. With the flask placed in its “permanent” location, the stopper should be quickly flipped upside-down on top of the flask so the flask remains sealed. The flask should be kept open the minimum amount of time possible. The T-pin should be used to attach the top (non-pigment end) of the paper to the center bottom of the rubber stopper.
  8. stopper and chromatogram into flask
    stopper into flask
    solvent front below the line
    solvent front below the line
  9. When the paper is securely attached, the stopper should quickly be flipped right-side-up and inserted into the flask in such a way that the paper does not touch the sides. The bottom of the paper should be barely in the solvent so that the solvent will be soaked up. It is imperative that you not move, jostle, or slosh the flask once the paper is soaking!
  10. solvent front above the line
    solvent front above the line
    done: solvent front by T end of pin
    done: solvent front by
    “bottom” end of pin
  11. As the solution is absorbed into the paper by capillary action, it will carry the various pigments up from the “center”. When the farthest band is about 1.0 to 0.5 cm away from the top of the paper or close to touching the T-pin, the chromatography may be stopped by removing the paper from the flask and replacing the stopper. The chromatogram should be observed and drawn, especially noting the colors of the various bands that are visible. The paper should be handled carefully, and no marks should be made on it.
  12. chromatogram removed and drying
    chromatogram removed and drying
    finished chromatogram
    finished chromatogram ready to cut apart

    Cutting Paper
  13. As a class, the identical bands will be put together and the pigments re-dissolved. One labeled 10 × 130 test tube will be supplied for each band Using (CLEAN) scissors, the various bands should be cut apart from each other (remember which is which). Each should then be placed as far into the bottom of the designated 10 × 130 test tube as possible. Everyone’s identical bands (i. e. all outer yellow bands) should go into the same tube to make the solutions as concentrated as possible. After everyone’s bands have been collected, the instructor (or an appointed class member) should place about 5 mL of 100% ethanol into each tube. Each tube should be labeled (if not done previously) and covered with Parafilm®. Each label should include the order and color of the band that tube contains (for example, “outer yellow”). The covered tubes should be placed in the designated rack for storage until the next lab period.
  14. redissolved pigments
    redissolved pigments
  15. All tubes being saved must be properly labeled and covered, then placed into a rack and stored in an appropriate location until next period. Chromatography solvent should be returned to the reagent bottle for reuse. All glassware should be washed and placed in the racks to dry. All scraps of chromatography paper and spinach should be disposed of properly and any other general clean-up should be done.
  16. Fluorescence
  17. Once the pigments have been re-dissolved in ethanol, your instructor may use the UV light to examine the tubes to demonstrate how chlorophyll (and any of the other pigments?) fluoresces.

Part B — Spectra of Pigments

  1. For this part of the experiment, students will be working in groups, based on the number of spectrophotometers available and the number of students in the lab section. (Optionally, as a class, a drop of each color of food coloring may be diluted with 100% EtOH so these may also be tested.) Someone in the class may grind a piece of spinach leaf or some dried parsley with a mortar and pestle, then add 100% EtOH to extract the plant pigments. Then, a small test tube, rack, glass funnel, and a piece of circular filter paper should be obtained. The test tube should be placed into the rack and the funnel into the test tube. The paper should be folded in half, then in quarters (half of half) as demonstrated by the instructor, then inserted into the funnel. The newly-extracted pigment solution should be poured through the filter paper to remove any particles. If this solution is very dark green, it will need to be diluted with more ethanol (see below).
  2. The tubes containing the isolated bands from the chromatograms (and those containing the diluted food coloring) will be distributed among the groups of students so that each group should have at least one of the redissolved, isolated bands and “something else” (mixed spinach or parsley pigments and/or food coloring and/or methylene blue or riboflavin) to test.
  3. One (CLEAN – without methylene blue stains) cuvette should be obtained for each solution the group will be testing plus one for plain EtOH, making sure to match glass colors (types/brands of cuvettes). Each cuvette should first be tested for the presence of unwanted, left-over methylene blue by placing a small amount of 100% EtOH in it, swirling, and holding the tube against a white surface. If a cuvette needs to be cleaned, do not use water to rinse it because all the solutions we will be testing are dissolved in EtOH, and water could interfere with the readings. For this experiment, only EtOH should be used to clean out cuvettes. Because it is so difficult to remove markings from cuvettes, and considering the possibility that any marks on them may interfering with the readings that will be taken, it is better to just line them up in the test tube rack in a pre-determined order corresponding to the labeled test tubes from the pigments being tested. In the unlikely case that it is necessary to label the cuvettes, ONLY PENCIL SHOULD BE USED, lightly writing only on the white area provided. DO NOT USE WAX MARKER OR LAB PEN! The cuvette that will serve as the blank should have about 4 or 5 mL of 100% ethanol added to it. Later, each pigment solution to be tested will be poured directly from its test tube into its own cuvette.
  4. While the redissolved, individual bands are probably dilute enough, if you are testing a solution of freshly-extracted, mixed parsley or spinach pigments, that may be too concentrated and may need to be diluted so the readings for it are not off the scale. Remember, Beer’s Law says that, by diluting a sample, its absorbance (at all wavelengths) will decrease.
    The chlorophylls in the mixture cause it to have an absorbance peak (highest amount of light absorbed) near 425 nm, so we want to adjust the concentration of the solution so that the A425 is not over 1.00. To do that, first, the wavelength on the spectrophotometer should be set to 425 nm. The zero and blank (using EtOH so that the machine subtracts out readings for whatever light the solvent absorbs) should be adjusted. Then, the absorbance of the mixed pigment solution should be read, and if the A425 is greater than about 1.000, it is necessary to dilute the sample. Ethanol should be added to dilute the solution and decrease the A425 to no more than 1.000. While it is possible to just “play around” with adding more ethanol or pigment solution until the absorbance is acceptable, if great accuracy is desired, the amount of alcohol needed can be calculated as follows.
    Recall that Beer’s Law says that, for example, if the absorbance reading is 2.00, theoretically, the addition of an equal volume of EtOH should dilute the sample to half its original concentration, such that the absorbance is half of the original, or only 1.00. Assuming you’re starting with 4 mL of pigment solution, the amount that you might typically use in a cuvette, Beer’s Law would mean that,
    A425 = 2.00  =  A425 = 1.00
    (x amt)/ ~4 mL soln(x amt)/ ~4 mL soln + ~4 mL EtOH
    where “(x amt)/# of mL” is an expression of concentration. The units used to express “x” could be moles, grams, or whatever, and since it’s the same on both sides, it cancels out and is not even necessary to know. This equation can be rewritten in general terms as,
    Ai (A425 observed)  =  Af (A425 desired of 1.00)
    (x amt)/ Vi (initial mL of soln)(x amt)/ Vf (initial mL of soln + mL of EtOH needed)
    which can be simplified to,
    Ai (A425 observed) × Vi (init mL of soln)   =     
          Af (A425 desired of 1.00) × Vf (initial mL of soln + mL of EtOH needed)
    and may be used to determine the amount of EtOH needed. When solved for milliliters of ethanol needed, this equation becomes:
    mL of EtOH needed =  Vi (initial mL of soln) × [Ai (A425 observed) – Af (A425 desired of 1.00)]
    Af (A425 desired of 1.00)
    If the desired, final absorbance reading is 1.00, this can be simplified to,
    mL of EtOH needed = Vi (initial mL of soln) × [Ai (A425 observed) – 1.00]
    Whether the amount of EtOH needed is calculated as just explained or whether a “guesstimate” amount is used, approximately the needed amount of alcohol should be added and a new A425 reading obtained. As needed, the volume should be adjusted further and another reading taken. When the A425 is 1.000 or slightly less, the concentration has been properly adjusted. About 4 to 5 mL of the diluted solution should be kept in (or placed in) a cuvette for testing.
  5. The isolated pigment bands are dilute enough that they are OK as is and do not need to be diluted. Each should be decanted into a separate, clean (check first for methylene blue) cuvette, taking care to not include any of the paper pieces.
  6. Absorbance readings should be obtained for all pigments at 25-nm intervals, and the process of obtaining readings will be quicker if all samples for which the group is responsible are tested at a given wavelength before changing to the next wavelength (all tested at 350 nm, then all at 375 nm, etc.). It is much more time-consuming to test one sample at all wavelengths, then go back and “start over” to test a second sample, etc. Initially, the wavelength should be set to 350 nm and the zero and blank rechecked. Absorbance readings should be obtained for all specimens the group is testing. setting the wavelength Then the spectrophotometer should be set at 375 nm, the zero and blank readjusted, and another set of readings obtained. Readings of absorbance should be taken at 25-nm intervals from 350 to 800 nm (350, 375, 400, etc.). Each time the wavelength is changed, it is necessary to recheck both the zero and the blank to get correct readings. Readings should be obtained for each of the bands being tested before changing wavelength. Readings should be recorded in students’ lab notebooks in chart form with columns for wavelength and for each of the samples. Also, data should be entered online.

Data

Part A — Paper Chromatography

The resulting bands on the chromatography paper should be drawn (then colored with colored pencils?) and described (color, location with respect to the solvent front and/or original spot). A tentative identification should be assigned to each of the pigments based on the list of pigment colors mentioned in the Background. From the colors of the individual bands on the chromatogram, which pigment does each of these bands appear to represent? Which is the smallest or fastest-moving molecule? Which is the slowest? Remember to draw any new equipment used.

Part B — Spectra of Pigments

  1. All spectrophotometer readings should be recorded in group members’ lab notebooks. A suggested format is:
    WavelengthPigment #1 (Name?)Pigment #2 (Name?)
    350 mnabsorbance?absorbance?
    375 mnabsorbance?absorbance?
    etc.
  2. Data for the absorption spectra of all solutions/bands tested should also be entered online (once per group — per set of data, not multiple entries of the same data). When all data have been entered, you may then return to the Web site to print out the class data.
  3. Graph of Data

  4. For each sample the group tested, a graph of wavelength (on the X- or horizontal axis) versus absorbance (on the Y- or vertical axis) should be constructed. The graphing protocol should be used as a reference on proper graphing techniques. Because this graph represents data which do not exhibit a proportional correlation, sequential points should be connected in “dot-to-dot” fashion, and the graph will not be a straight line graph. Absorption maxima (peaks) and minima for each of the solutions tested should be noted.
    The example, above, is a graph of the spectra for two concentrations of Chlorophyll A, represented by the black line and the greenish line, and the spectrum for Carotene, represented by the pinkish line. Because the concentrations of the solutions were not standardized in any way, the heights of the peaks (which, you should recall, are merely concentration-dependent) are not significant (differences in concentrations of solutions are not being examined in this experiment), but rather, as notated in the example, above, the locations of the peaks, the maxima, as well as the minima, relative to the wavelengths tested, are important data. Thus it is important that Chlorophyll A’s maximum is at 425 nm as compared to Carotene’s maximum at 450 nm, and it is important that Chlorophyll A’s minimum is at 525 nm as compared to Carotene’s minimum at 600 nm. For this experiment, we don’t care that at 425 nm, the absorbance for one of the Chlorophyll A solutions was 0.59 and the other was 0.29 — all that means is that one solution was about twice as concentrated than the other, but the important thing is that Chlorophyll A had a maximum absorbance peak at 425 nm. It is also important that Chlorophyll A has a second maximum at 675 nm.
    Also, because these glass cuvettes are only good in the visible range (UV takes special quartz cuvettes and IR takes special salt cuvettes), the “drift” at the beginnings and ends of the graph, where the wavelength is approaching the ultraviolet or infrared range are “meaningless” for this experiment.
  5. At what wavelength(s) did each of the isolated pigments absorb the most/least light? Do the observed absorption maxima and minima correspond to those reported in the literature for each of those pigments? Were the tentative identifications of the bands correct — do the absorption data support the identifications made based on color/appearance? To what colors do these wavelengths correspond?
  6. The absorption spectrum of the mixed pigments tested should be compared with the spectra from the various “known” pigments. By matching the peaks, which of the individual pigments does the mixed pigment solution contain? Also, at which wavelengths did the mixed plant pigments absorb the most light — where were the absorbance peaks? To what colors do these wavelengths correspond? At which wavelength did they absorb the least light — where was the absorbance closest to zero? To what color does this correspond?
  7. If methylene blue, food coloring, or any other pigments were also tested, the same analysis should be done for each pigment tested. The absorption maxima and minima should be determined for each of the colors tested. What wavelength(s) of light is/are each of the colors absorbing (therefore unavailable to a plant), and what wavelength(s) is/are each color not absorbing (therefore reflecting or transmitting and available to a plant). If a plant was placed into a solution containing this/these pigment(s), what wavelength(s)/color(s) of light would be available to the plant to use?
  8. Any other significant notes, observations, and data should be included.

Optional Additional Experiment(s)


Things to Include in Your Notebook

Make sure you have all of the following in your lab notebook:


Copyright © 2011 by J. Stein Carter. All rights reserved.
Based on printed protocol Copyright © 1986 D. B. Fankhauser
and © 1989 J. L. Stein Carter.
Chickadee photograph Copyright © by David B. Fankhauser
This page has been accessed Counter times since 5 Sep 2011.