SPECTROSCOPIC ANALYSIS OF...

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Spectroscopic Analysis of Chlorophyll Revised: 12/13/14 1 SPECTROSCOPIC ANALYSIS OF CHLOROPHYLL Adapted from the Journal of Chemical Education Vol. 88, No. 8, May 2011 Vol. 81, No. 3, March 2004 INTRODUCTION Plant pigments collect light from the sun which, when combined with carbon dioxide and water, convert that energy into the chemical bonds of carbohydrates (the starches and sugars that provide the building blocks for plant growth). Figure 1. Plant Respiration Many pigments are present in a green plant leaf including (but not limited to): chlorophyll, carotene, and xanthophyll. Chlorophyll is the most dominate of these pigments, absorbing specific wavelengths of the visible light when collecting energy from the sun. The wavelengths not absorbed are transmitted to our eyes, leading to the observation of green colored leaves. Accessory pigments (like carotene and xanthophyll) absorb at other wavelengths and/or participate in the electron transport chain, helping the chlorophyll in its conversion of light energy to chemical energy. A chlorophyll molecule is made of two distinct parts. The chlorophyll molecule’s tetrapyrrolic porphyrin ring acts as a chelate, binding a magnesium ion in a rigid square planar arrangement. The Mg ion, in conjunction with the ring, is capable of a variety of oxidation states allowing for the acceptance or donation of electrons as part as the conversion of light energy into plant growth (the electron transport chain). This magnesium porphyrin head is connected by an ester functionality to a hydrophobic chain (tail) derived from a phytol. This tail anchors the chlorophyll molecule into the lipid membranes of the chloroplast. bioweb.uwlax.edu

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  • Spectroscopic Analysis of Chlorophyll Revised: 12/13/14

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    SPECTROSCOPIC ANALYSIS OF CHLOROPHYLL

    Adapted from the Journal of Chemical Education Vol. 88, No. 8, May 2011

    Vol. 81, No. 3, March 2004

    INTRODUCTION Plant pigments collect light from the sun which, when combined with carbon dioxide and water, convert that energy into the chemical bonds of carbohydrates (the starches and sugars that provide the building blocks for plant growth).

    Figure 1. Plant Respiration Many pigments are present in a green plant leaf including (but not limited to): chlorophyll, carotene, and xanthophyll. Chlorophyll is the most dominate of these pigments, absorbing specific wavelengths of the visible light when collecting energy from the sun. The wavelengths not absorbed are transmitted to our eyes, leading to the observation of green colored leaves. Accessory pigments (like carotene and xanthophyll) absorb at other wavelengths and/or participate in the electron transport chain, helping the chlorophyll in its conversion of light energy to chemical energy. A chlorophyll molecule is made of two distinct parts. The chlorophyll molecules tetrapyrrolic porphyrin ring acts as a chelate, binding a magnesium ion in a rigid square planar arrangement. The Mg ion, in conjunction with the ring, is capable of a variety of oxidation states allowing for the acceptance or donation of electrons as part as the conversion of light energy into plant growth (the electron transport chain). This magnesium porphyrin head is connected by an ester functionality to a hydrophobic chain (tail) derived from a phytol. This tail anchors the chlorophyll molecule into the lipid membranes of the chloroplast.

    bioweb.uwlax.edu

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    Porphyrin Head

    Phytol Tail

    Ester functionality joining Head & Tail

    tetrapyrrolic: containing 4 pyrrole rings

    Figure 2. Structure of Chlorophyll a

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    A few different forms of chlorophyll exist. Chlorophyll a & b are the two most common - differing only by a substituent attached to the pyrrole ring on the porphyrin ring opposite the phytol tail. On chlorophyll a the substituent is a simple methyl (-CH3) group; chlorophyll b has an aldehyde functionality (-C(=O)H) at the same location. Figure 3. Porphyrin Head of Chlorophyll b

    Pheophytin is a demetallated form of chlorophyll. The magnesium ion is replaced by two hydrogen atoms one to each of the negatively charged nitrogens in Figure 3 above. Pheophytins are part of the electron transport chain and also form as chlorophyll degrades (resulting in the darkening and browning of vegetables upon prolonged cooking). Carotenoids are completely hydrophobic, existing within the lipid membranes. These accessory pigments absorb light energy and transfer that energy to chlorophyll. However, their most

    Figure 4. Carotenoids important role is protecting plants from free radicals (unpaired electrons) formed by ultraviolet radiation. The two most common carotenoids are carotenes and xanthophylls (see Figure 4).

    More polar aldehyde functional group

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    All of these pigments are brightly colored pigments indicating they readily absorb visible and some ultraviolet radiation. In this experiment, the absorbance spectra (A vs. (nm)) of 4 different solutions will be measured: spinach leaf extract, pure chlorophyll, yellow food dye, and blue food dye. As seen in previous experiments using absorbance spectroscopy, the color (wavelength, max) of light a compound absorbs is complementary to the color observed. While the simple spectra of the food dyes demonstrate this correlation, chlorophyll does not. Furthermore, the spinach leaf extract, being a mixture of the aforementioned pigments, is even more complex. Figure 5. Absorbance (Electron Excitation). As discussed in previous experiments and in extra assigned reading, the absorbance of a particular

    wavelength of light by a chemical is the result electron excitation (Figure 5). How does the electron release energy and get back to the ground state? The most common pathway is the transformation of the energy into vibrational motions of the molecule (heat). However, for some molecules light is emitted when the electron returns to the ground state. This phenomenon is called fluorescence.

    Figure 6. Fluorescence (Radiative Emission). Fluorometers are the instruments used to measure fluorescence. Like visible spectrometers, they have a light source, a sample holder, and a detector. The source exposes the sample to a certain wavelength of light and the detector measures the light coming out of the sample. For a visible

    cuvette holder or cuvette

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    spectrometer, the source, sample, and detector are arranged in a straight line, so the source light transmitted (not absorbed by) the sample is measured by the detector. For a fluorometer, the source, sample, and detector are arranged in a right angle, so the detector only measures the light emitted from the sample. In lab, you will use a spectrometer (SpectroVisPlus, Vernier) capable of measuring both fluorescence and absorbance. Figure 7a. Spectrometer Figure 7b. Arrangement of source, sample, & detector. The maximum wavelength (max) of absorption must be known to acquire a fluorescence spectrum. In an analytical grade instrument, the sample is excited at max, and the resulting emission is recorded at longer wavelengths (lower energy) (Figure 8). (When using the SpectroVis Plus spectrometer, two excitation wavelengths are available: 405 nm and 500 nm. Choose the wavelength closest to max.) This lowering of energy from absorbance to fluorescence Figure 8. Overlay of Absorbance & Fluorescence Spectra. measurements is referred to as the Stokes Shift. An electron is excited into a higher rotational

    level of the LUMO electronic level. The electron is able to relax to the LUMOs lowest rotational level before returning to the HOMO and emitting a photon. Because of the relaxation, the energy gap is smaller, therefore, the photon released has less energy (and a longer wavelength). (Figure 9.)

    Figure 9. Rotational & Electronic Energy Levels Like absorbance, fluorescence has a linear relationship with concentration, as long as the solution is

    dilute. In Beers Law, absorbance is equal to solution concentration multiplied by path length (the

    distance light travels through solution) and molar absorptivity (a constant unique to each solute (A =

    cl). The calculation of fluorescence intensity is a bit more complex, as seen below:

    vernier.com

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    F = kQPo(2.303 cl)

    F is the measured fluorescence intensity, k is a geometric instrumental factor, Q is the quantum

    efficiency (photons emitted/photons absorbed), Po is the radiant power of the excitation source. Just like Beers Law, is the molar absorptivity coefficient, c is the concentration, and l is the path

    length. For our purposes, the equation can be simplified to express the direct, linear relationship

    between fluorescence intensity and concentration:

    F = Kc

    During the first week of this experiment, absorbance measurements, in conjunction with Beers Law, will be used to determine molar absorptivity of chlorophyll by measuring the absorbance of a standard solution of chlorophyll. This value will allow for the calculation of chlorophyll concentration in the spinach extract. The chlorophyll concentration in the extract will also be found with fluorescence. Fluorescence spectra will be acquired for standard chlorophyll solutions to create a calibration curve similar to a Beers Law Plot. During the second week, spinach will be extracted, analyzed by thin layer chromatography (TLC), and its chlorophyll content will be quantified by comparing the absorbance and fluorescence spectra to the work done in week 1.

    SAFETY PRECAUTIONS

    Safety goggles, aprons, and gloves must be worn at all times in the laboratory. Heptane and acetone

    are extremely flammable and harmful by inhalation, ingestion, and when in contact with skin. Any

    container holding either heptane or acetone should be capped when not in use to prevent

    evaporation of the solvents, as they are harmful when inhaled. Heptane and acetone solutions must

    be placed in appropriate waste bottles and can NEVER be poured down the drain. Report all spills,

    accidents, or injuries to your TA.

    Before starting the experiment, the TA will asks you to do a quick demonstration or talk-through one of the following: 1) How to prepare a TLC plate and a TLC developing chamber? 2) When do you need to remove the TLC plate from the developing chamber and how do you calculate Rf? 3) How to set up a vacuum filtration? 4) How to use a separatory funnel, specifically: How and when do you vent a separatory funnel and how do you drain liquid from the separatory funnel? 5) How to use a separatory funnel, specifically: How and when do you vent a separatory funnel? 6) How to use a separatory funnel, specifically: How do you drain liquid from the separatory funnel?

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    Make sure you watch the videos on the course website and read the documents to prepare. These demonstrations will be done every week. Everyone will have presented at least one topic by the end of the quarter. The demonstrations should be short (>1 min) and will be graded. General Experiment Information:

    Work in pairs for all parts. Answer italicized questions imbedded in the procedures in your observations. Discuss the questions with your TA. (You need to be actively thinking about the answers, the TA is only a guide.) Attach spectra (LabQuest2 file and screenshot) to your ELN. Label all peaks with wavelength and absorbance values.

    WEEK 1

    Part A. Measuring Absorbance of Food Coloring and Chlorophyll Standard

    1. Connect the SpectroVis Plus to the LabQuest2, and calibrate the spectrometer:

    a. In the screen, click the box labeled USB:Abs. Select calibrate from the drop down menu. (Allow the lamp to warm up).

    b. With a ruler, measure the path length (inner width) of the cuvette.

    c. Prepare a blank by filling an empty cuvette with deionized water. Why is deionized water used as the blank?

    d. Wipe the outside of the cuvette with a Kimwipe and place the blank in the spectrometer. When taking an absorption measurement, always align the cuvette with the same face (mark the top of the cuvette with a grease pencil) pointing toward the detector. Why is this necessary? Once the warm-up period is complete, select Finish Calibration and OK.

    Absorbance Spectra of Food Coloring

    2. Collect an absorption spectrum of blue food coloring solution:

    a) Empty the blank cuvette, prerinse twice with small amounts of the blue food coloring

    solution, then fill with the blue liquid. Always prerinse a cuvette with small amounts of the

    solution that it will contain. Why?

    b) Place the cuvette in the spectrometer and in the screen, click . A spectrum will be

    displayed. Click . Examine the graph, noting peak(s) of high absorbance and other

    distinguishing features.

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    3. Repeat with the yellow food coloring solution and a 1:1 mix of the blue and yellow solutions.

    Store the run by selecting the store option after you hit again. You do not need to

    recalibrate the spectrometer in between the three absorption spectra collected above. Why?

    4. To view all runs overlapped, click on the box that says Run 3 and select All Runs. Send this

    data to your ELN.

    Absorbance Spectrum of Chlorophyll Stock Solution

    5. Choose New from the File menu. Prepare a blank by filling an empty cuvette with heptane.

    Place the blank in the spectrometer. Select Finish Calibration and OK. Why is heptane used

    as the blank? Why did the spectrometer need to be recalibrated?

    6. From your TA, obtain a vial filled with 6.0 ppm chlorophyll solution. This is the stock solution

    and heptane is the solvent. This solution was prepared and then placed in the freezer by the

    stockroom. You must wait until the solution has reached room temperature before continuing.

    Pour this solution into a cuvette, insert into the spectrometer, and click . Click . Do

    not remove the chlorophyll stock solution from the spectrometer. Examine the graph, recording

    the and A of the peak(s) in the spectrum. Send this data to your ELN. What is the molarity of

    a 6.0 ppm solution?

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    Part B. Fluorescence Measurement of Chlorophyll Standard

    1. Go to the screen, click on the USB: abs box and from the dropdown menu select Change

    Units Fluorescence (405 or 500 nm). Choose the correct wavelength (405 or 500 nm) based

    on your evaluation of the absorption spectrum just taken. Explain your wavelength choice.

    2. Choose Sensors > Data Collection. Change the sample time to 100 ms. Click OK. In the

    screen, click . The stock solutions fluorescence spectrum will be displayed. Click .

    Send this data to your ELN.

    3. In the screen, click Sensors > Data Collection. Mode: Event w/ Entry, Name:

    Concentration, Units: ppm. Do not change the wavelength setting. What is the ? Why has the

    spectrometer chosen that wavelength? Click OK to continue.

    4. Click . Once the reading stabilizes click Keep. Enter the concentration (in ppm) of your

    stock solution and click OK. This is the first standard (100% the original concentration) for the

    calibration curve you will create. Record the indicated data in the table below.

    Include this serial dilution in the observations of your ELN. Dilution % Original

    Concentration Concentration

    (in ppm) Fluorescence

    Intensity at :

    ______nm 0 100 (original) 6

    1 50

    2 25

    3 12.5

    4 6.25

    5. Dilution #1 (50% of original concentration): Add exactly 2 mL the chlorophyll standard

    solution to a clean, dry 10 mL graduated cylinder. Excess solution should be placed in the

    organic waste container in the hood. Add 2 mL of heptane to the graduated cylinder. Calculate

    the new concentration. Place the solution into the cuvette. After the reading stabilizes, click

    Keep. Enter the concentration and click OK. Record the indicated data in the table below.

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    6. Dilution #2 - 4: Repeat the dilution and fluorescence measurements until 4 dilutions have been

    made. When finished, click Stop to end data collection.

    7. Add a Linear Regression (r2 value) and Trendline Equation to your graph. What does the r2

    value represent?

    Make sure to clear your email address and password of the LabQuest2 so others cant access your email account. Shutdown the LabQuest2 and not simply put it to sleep. To shutdown the LabQuest2: press the home key, select System Shut Down OK.

    WEEK 2

    This week, various pigment molecules in spinach leaves will be extracted. The absorbance and

    fluorescence of the spinach extract will be measured and compared with last weeks results. Thin

    layer chromatography (TLC) will be performed to separate and identify the various pigments within

    the spinach leaf: carotenes, chlorophyll a and/or b, pheophytin, and xanthophylls.

    Part C. Spinach Leaf Extraction

    Note: Make sure you have all your glassware prepared BEFORE beginning the extraction!

    1. Remove the stems from ~3 g of spinach leaves, record their mass. Tear spinach leaves into tiny

    pieces and then, in the hood, use a mortar and pestle to grind the spinach leaves with 3 mL

    acetone (~2 minutes). (Acetone is a volatile organic liquid so you may need add more because

    of evaporation.) Why do the spinach leaves need to be ground?

    2. Add 10 mL DI water to the pulpy mixture, and pour this solution into a 50 mL flask. Then add

    10 mL heptane to the mortar and pestle to wash out any remaining spinach leaves into the flask.

    Swirl the mixture gently for ~1 minute.

    3. Vacuum filter the mixture to separate the extract from the leaves. Use the 125 mL Erlenmeyer

    collection flask. Make sure to use filter paper during the filtration! Pour the spinach extract

    from the flask, while trying to avoid pouring all of the spinach leaves into the filter (its fine if

    some are poured out). Keep filtering the extract until the spinach leaf pulp left on the filter

    paper looks relatively (but not completely) dry. If it looks like some pulp went through the filter

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    and into your collection flask, quickly filter the extract one more time, using a different piece of

    filter paper. Dispose of filter paper containing spinach leaves in the trashcan.

    4. Pour the filtrate into the separatory funnel, cap, then swirl gently, and vent the funnel three

    times. Let the separatory funnel sit for two minutes to allow the heptane and water to separate.

    Why does the sep funnel need to be vented?

    5. Shine a UV lamp on the separatory funnel. One of the layers fluoresces (the room needs to be

    dark, ask your TA to dim the lights). Which layer is organic? Which is aqueous? How do you

    know? Which layer fluoresces and why?

    6. If you have an emulsion, discuss the addition of a brine solution with your TA. If you do not

    see two layers, add 1-2 mL of heptane. Drain the aqueous layer into a beaker by opening the

    stopcock. (Do not discard this layer, in case you made a mistake.) Next, drain the organic layer

    out of the separatory funnel into a clean dry test tube labeled Extract #1.

    7. Add just enough anhydrous MgSO4 to Extract #1 to fill the bottom tip of the test tube (about 10-

    15% of the extracts total volume). Using a stirring rod, mix the contents of the test tube. What

    does the anhydrous MgSO4 do? How does it interact with the extract? What does it become?

    8. Use a disposable pipet to carefully separate the extract from the MgSO4. You can do this by

    decanting off the solution into another test tube, labeled Extract #2, without disturbing the

    solids. You do not need to decant all of the solution, just enough to do your analysis (~1-2 mL),

    but try to get as much as you can. After the transfer, place a cap on the Extract #2 tube. Now,

    place Extract #2 under the UV lamp. Try both the short and long wavelength setting. Which

    wavelength produces fluorescence? Why do you think this wavelength produces fluorescence

    while the other does not?

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    Part D. Thin Layer Chromatography of Spinach Leaf Extract

    1. Place a piece of 5.5 cm filter paper so that it touches the bottom of the TLC

    developing jar. Pour ~5 mL of a heptane:acetone solution into the jar (the

    liquid should fill the bottom of jar to a depth of ~0.5 cm) and close with a cap.

    The filter paper will become wet with the solvent and ensure the atmosphere

    inside the chamber is saturated with solvent vapor.

    2. Prepare the TLC plate by lightly drawing a line on the plate 1 cm from the

    bottom with a pencil. Use TLC spotters to transfer spinach extract drop by drop to the plate.

    Make the spots as small as possible (~1 mm diameter) to achieve optimal separation. If your

    spinach extract is relatively concentrated (creating a green spot almost as intense as the color of

    the spinach leaves just extracted), one drop on the TLC plate may be sufficient. If not, you may

    need to add more than one drop repeatedly to the same spot. Allow the plate to dry between

    each drop. Note: Dispose of TLC spotters in the Broken Glass containers only. Any type of

    glass in the trashcans is a health hazard for the janitorial staff.

    3. Let the TLC plate dry for 1 minute, then place in the TLC chamber and cap. The TLC plate

    should not touch the filter paper. (If the filter paper touches the side of the TLC plate, the spots

    will be drawn sideways.) Why does the TLC chamber need to be capped while the TLC plate is

    developing?

    4. Solvent will move up the plate wetting it. Once the solvent front is within ~0.5 cm of the

    top of the plate, remove the plate from the chamber and mark the solvent front line with a

    pencil. Do not let your solvent front travel to the top of the silica on the TLC plate, Rf values

    cannot be calculated if the solvent front is not visible.

    5. Circle visible spots on the TLC plate with a pencil. Use a UV lamp to see spots only visible in

    the UV (usually white or lightly colored compounds). Take a digital picture of the plate and

    attach it to your ELN. In a table, record the color and Rf values of each spot. Provide one

    sample Rf calculation, clearly indicating where the values came from on your TLC plate.

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    Part E. Measuring Absorbance of Spinach Leaf Extract

    Reminder: Send all files to your ELN.

    1. Connect the SpectroVis Plus to the LabQuest2, and calibrate the spectrometer: In the

    screen, click the box labeled USB:Abs. Select calibrate from the drop down menu. (Allow the

    lamp to warm up). Prepare a blank by filling an empty cuvette with heptane. Wipe the cuvette

    off with a Kimwipe. Place the blank in the spectrometer. Once the warm-up period is complete,

    select Finish Calibration and OK.

    2. Measure the absorbance spectrum of the spinach leaf extract. Depending on how well the

    extraction went, your solution might need to be diluted. The goal is to systematically create a

    solution with an absorbance similar to the chlorophyll a standard from Part A. Ask the TA for

    some guidance to avoid overdiluting. A large dilution may be required initially, followed by

    serial dilutions (as in Part B). Be sure to record the steps and volumes used in the dilution

    process so that you can calculate the original chlorophyll concentration in Extract #2.

    Part F. Measure the Fluorescence of the Chlorophyll Extract

    1. Go to the screen, click on the USB: Abs box and from the dropdown menu select Change

    Units Fluorescence (405 or 500 nm). Choose the wavelength based on last weeks max.

    2. Choose Sensors > Data Collection. Change the sample time to 100 ms.

    3. Obtain the fluorescence spectrum of the solution created in Part E. Click in the

    screen. The fluorescence spectrum of the extract will be displayed. Click . Send this

    spectrum to your ELN.

    4. Compare the fluorescence intensity at max with the calibration curve created week 1. If the

    value is above the highest fluorescence intensity of that curve, continue the serial dilution

    (started in Part E above) until the fluorescence intensity falls between the upper and lower limits

    of the calibration curve. Continue to record the values in the table started in Part E they will

    be needed to find the concentration of the undiluted spinach leaf extract.

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    Make sure to clear your email address and password of the LabQuest2 so others cant access your email account. Shutdown the LabQuest2 and not simply put it to sleep. To shutdown the LabQuest2: press the home key, select System Shut Down OK.

    DISCUSSION Spectra or figures should be incorporated seamlessly into one document with the answers.

    Combining 2 or more spectra in a single plot may be useful and no spectra or figure should take up

    an entire page.

    1. In this experiment, you took the absorption spectra of 5 different solutions: yellow food dye,

    blue food dye, a mixture of yellow and blue food dye, chlorophyll a standard, and spinach leaf

    extract. The spectrum of chlorophyll b is available on the course website. Compare and

    contrast these absorption spectra. (All spectra must be shown, and all should have x and y axes

    with the same minima and maxima.)

    Connect the max value(s) with solution color. What is the effect when a compounds

    spectrum has more than one max?

    If you were given these five spectra without any title or other identifiable information and

    then separately given a list with the identities of the four solutions, how would you go about

    matching solution identity to each spectra? Provide the scientific reasoning used.

    Does solvent matter? Why or why not?

    2. What is the chemical identity of the spots on your the TLC plate? (Include a labeled image of

    the TLC plate.) Explain the reasoning behind your assignments use fundamental chemical

    knowledge (polarity and observed color, for example). List any references used.

    3. Using the absorbance spectra chlorophyll a, and chlorophyll b, and the color of the other

    pigments (from the TLC), indicate where the pigments are absorbing on the extracts spectrum.

    4. What was the excitation wavelength used to create the fluorescence spectra in this experiment?

    Why was that wavelength used?

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    5. Using data from week 1 and 2, determine the concentration (in ppm) of chlorophyll a in the

    spinach leaf extract from the absorbance measurements. Chlorophyll has two max values,

    calculate the values below at both.

    Use Beers Law to find the molar absorptivity () of chlorophyll a from week 1s data.

    Using this calculated value of just found, find the chlorophyll a concentration (in ppm) of

    the diluted spinach extract solution.

    What is the chlorophyll a concentration (in ppm) in the undiluted spinach extract?

    6. Using data from week 1 and 2, determine the concentration (in ppm) of chlorophyll a in the

    spinach leaf extract from the fluorescence measurements.

    Use the Beers Law type plot from week 1 to find the concentration of chlorophyll a or b

    in your spinach extract. Using this plot, find the concentration of chlorophyll a in the

    diluted spinach extract solution.

    What is the chlorophyll a concentration (in ppm) in the undiluted spinach extract?

    7. Why might the concentration calculated in #5 be different from #6?