Materials Included In Kit

Additional Materials Required

Prelab Preparation

1. Slurry of each of the extract powders.
   1. Pour each of the extract powders into separate, labeled 50-mL beakers.
   2. Add 8 mL of acetone to each beaker.
   3. Swirl for several minutes.
   4. Cover to prevent evaporation of the acetone.
2. All materials and solution used in the preparation of the chloroplast suspension should be refrigerated overnight and kept cold prior to making the suspension. This includes the pitcher of the blender, the beakers, the funnel and the phosphate solution.
3. Phosphate buffer solution. Prepare at least one day prior to making the chloroplast suspension.
   1. Dilute 50 mL of the 10x concentrated phosphate buffer to 500 mL with deionized water.
   2. Cover and store on ice or in a refrigerator until ready to use.
   3. The solution is stable for four weeks if kept cold.
4. Chloroplast suspension. Prepare the chloroplast suspension as close to lab time as possible (within the same day).
   1. Remove the ribs from the fresh baby spinach leaves.
      
   2. Place approximately 3 g of the fresh baby spinach leaves behind a heat sink (large beaker filled with tap water) in front of a strong light source for 1–3 hours prior to preparing the chloroplast suspension.
   3. In a blender, add 3 g of the baby spinach leaves to 300 mL of cold phosphate buffer solution.
   4. Blend for 30 to 40 quick pulses followed by a 30 second blend.
   5. Place a 400-mL beaker in an ice bath. Place four layers of cheesecloth into a large funnel, and place the funnel into the beaker.
   6. Filter the suspension through the cheesecloth in the funnel and into the beaker. Squeeze the cheesecloth to retrieve as much buffer as possible. Note: You should be able to retain 90% of the original volume (≈270 mL).
   7. Pour the chloroplast suspension into an amber bottle or completely cover the beaker with aluminum foil. Label the container “active chloroplast suspension.”
   8. Store on ice or in a refrigerator until ready to use. Note: During the experiment, the active chloroplast suspension must be kept on an ice bath and in the dark prior to being incubated by the students.
   9. The success of this laboratory hinges upon the preparation of the chloroplast suspension.
   1. Too concentrated or too weak of a suspension will adversely affect the results. Test the suspension prior to the lab. If the DCPIP is reduced in the unheated sample within 5 minutes of light exposure, dilute the chloroplast suspension with additional buffer or add fewer milliliters of suspension to each test tube. Note: Even adding as few as 5 drops is possible if the chloroplast solution is very concentrated.
   2. If the DCPIP is not significantly reduced within 15 minutes, increase the amount of chloroplast suspension added or decrease the concentration of the DCPIP solution.
   3. (Optional) Test the activity of the chloroplast suspension. Set a warmed up spectrophotometer at 600 nm. Zero the spectrophotometer with 5 mL of the phosphate buffer. Insert a cuvet containing 5 mL of the new chloroplast suspension. An ideal absorbance reading is 0.300–0.400.
5. Heated chloroplast suspension.
   1. Place 50-mL of the active chloroplast suspension into a 100-mL beaker.
   2. Place in a 50 °C hot water bath for 5 minutes. Note: Do not boil the suspension as it will clump. If it clumps, use a glass stirring rod to break the clumps apart.
   3. Pour the heated chloroplast suspension into an amber bottle or completely cover the beaker with aluminum foil. Label the container “heated chloroplast suspension.”
   4. Store on ice or in a refrigerator until ready to use. Note: During the experiment, the heated chloroplast suspension must be kept on an ice bath and in the dark prior to being incubated by the students.
6. 2,6-Dichlorophenolindophenol (DCPIP) solution, 0.1% (also known as 2,6-Dichloroindophenol, DPIP and DCIP).
   1. Using a magnetic stirrer and stir bar, dissolve 0.1 g of DCPIP in 100 mL of deionized water. Note: Slightly lowering the pH of the solution enables the DCPIP to more easily dissolve.
   2. Transfer the solution to an amber bottle with cap or completely cover the beaker in aluminum foil.
   3. Store on ice or in a refrigerator until ready to use. Note: During the experiment the DCPIP solution must be kept on an ice bath and in the dark prior to being incubated.
   4. The solution is stable for two weeks if kept cold.
7. Incubation area. Use one of the three methods described below.
   1. Place a 1-L beaker filled with tap water between a 150 W flood lamp and the area in which students will incubate their samples. The water serves as a heat sink for the light so that the chloroplast suspensions do not become heated.
   2. Turn on a bright overhead projector and point the light directly at the area in which students will incubate their samples. The overhead projector should be located close to the incubation area to ensure a very bright light source.
   3. Turn on a bright overhead projector and place the cuvets in a beaker filled with water on the transparency area of the overhead projector (see Figure 6 in the Procedure).

Safety Precautions

Acetone and the chromatography solvent are dangerous fire risks; flammable; toxic by ingestion and inhalation. This lab should be performed only in an operating chemical fume hood or well-ventilated area. Wear chemical splash goggles, chemical-resistant gloves and a chemical-resistant apron. Remind students to wash hands thoroughly with soap and water before leaving the laboratory. Please consult current Safety Data Sheets for additional safety, handling and disposal information.

Disposal

Please consult your current Flinn Scientific Catalog/Reference Manual for general guidelines and specific procedures, and review all federal, state and local regulations that may apply, before proceeding. 2,6-Dichlorophenolindophenol may be disposed of according to Flinn Suggested Disposal Method #26b. Acetone, chromatography solvent, spinach extract and blue-green algae extract may be disposed of according to Flinn Suggested Disposal Method #18a. Phosphate buffer may be disposed of according to Flinn Suggested Disposal Method #26b. TLC plates may be disposed of according to Flinn Suggested Disposal Method #26a.

Lab Hints

• Enough materials are provided in this kit for 8 groups of students. Both parts of this laboratory activity can reasonably be completed in two 50-minute class periods. Any prelaboratory assignment should be completed before coming to lab, and the data compilation and calculations can be completed the day after the lab.
• Good technique is important to achieving clean separations in thin-layer chromatography. Common sources of student error include “overloading” the TLC plates by placing too much extract on the initial spot and band broadening that occurs because the initial spot is too large. Another common problem is “staining” that occurs when the pigment spots are submerged below the solvent level.
• Ensure students label all materials with their group number and the activity number or chemical used in the container.
• The chromatography solvent contains 90% petroleum ether and 10% acetone. Do not discard leftover chromatography solvent—the solvent may be recycled. Save it for use by another class or for another chromatography experiment. Do not leave the chromatography solvent uncovered for long periods of time—one component may evaporate faster than the other, changing the polarity of the mixture.
• Activity 2 may be conducted in a dim room or in a closed drawer or cupboard to control the amount of light exposure each cuvet receives prior to its incubation in the bright light.
• Percent transmittance was chosen as the units for Activity 2 so that students would not confuse absorbance in a spectrophotometer with adsorption in chromatography.
• If there is a limited number of spectrophotometers, assign two cuvets to each group and pool the class data.
• Pool the class’s data for Activity 2 and discuss sources of variation and error, such as instrument drift, error in cuvet alignment, measurement errors and instrument precision.

Teacher Tips

• Many plant pigments fluoresce under ultraviolet light. Once students’ TLC plates have dried, shine a UV light source very close to the plate—you should see many or all of the pigments fluoresce! Note: A black-light bulb will not work.
• Extend Activity 1 by having students prepare extracts from other plants or algae. Tear and grind the leaves with a mortar and pestle and a small amount of clean sand and acetone to extract the pigments. Additional chemicals and materials are available from Flinn Scientific, Inc.
• Extend Activity 2 by using colored filters to block specific wavelengths of light from reaching the active chloroplasts or by testing the rate of photosynthesis at different temperatures or different light intensities.
• Adsorption refers specifically to the adhesion of atoms, ions or molecules onto the surface of another substance. Absorption refers to the penetration of one substance into the inner structure of another.

Sample Data

Activity 1

Observations

Blue-Green Algae Activity 2

Observations and Analysis  Graph the data for the reduction of DCPIP by light. For this graph you will need to determine the following:

1. What is the dependent variable? Percent Transmittance
2. What is the independent variable? Time in minutes
   {10779_Answers_Figure_7}

Answers to Questions

Activity 1

1. What factors are involved in the separation of the pigments?

   The polarity of the pigments varies because of differences in the structures of the pigment molecules. The different polarities cause the pigments to adsorb differently on the TLC plates. The polarity of the solvent and the affinity for the adsorptive surface affect the separation of the pigments. The time and distance allowed for the separation to occur also affect the separation of the pigments.

2. Would you expect the R_f value of a pigment to be the same if a different solvent were used? Explain.

   No, the R_f value will vary with different solvents. The R_f values will change due to variations in time, distance traveled, type of adsorptive surface used with a particular solvent also causes the solvent front move at a different rate.

3. Which pigment directly captures light energy? What are the roles of the other pigments?

   Chlorophyll a is the only pigment that is able to directly capture energy from light. The other pigments also capture light, but that energy must be transferred to chlorophyll a before ATP can be generated and NADP is reduced.

Activity 2

1. What is the purpose of DCPIP in this experiment?

   DCPIP replaces NADP as the acceptor of the electron removed from water as the result of the photo-oxidation of chlorophyll a.

2. What is the purpose of the following cuvets: the zero cuvet, the blank cuvet, and the control cuvet?

   The zero cuvet serves as the instrument blank. It ensures that the baseline of the spectrophotometer does not drift.

   The blank cuvet serves as a reference for the amount of light absorbed by the suspension of chloroplasts. The readings should remain approximately the same for the entire experiment.

   The control cuvet serves as a reference to the color of oxidized DCPIP since there is no chloroplast to facilitate its reduction to colorless. The readings should remain approximately the same for the entire experiment.

3. What was measured with the spectrophotometer in this experiment?

   The spectrophotometer measures the amount of light that is transmitted through a solution of DCPIP and chloroplasts at 600 nm.

4. What reasons can you give for the difference in the rate of photosynthesis between the active chloroplasts that were incubated in the light and those that were heated?

   The heat caused the chloroplasts to become inactive. The chlorophyll a is no longer able to capture an electron from the water and reduce the DCPIP.

5. What reasons can you give for the difference in the rate of photosynthesis between the active chloroplasts that were incubated in the light and those that were kept in the dark?

   Photosynthesis is occurring in the active chloroplasts therefore the DCPIP is being reduced by electrons flowing in the photosynthetic electron transport chain.

References

Biology: Lab Manual; College Entrance Examination Board: 2001.

Introduction

In plants, algae and some types of bacteria, photosynthesis is the process that traps the energy from sunlight, called photons, to convert carbon dioxide and water to glucose and to make adenosine triphosphate (ATP). ATP is the “fuel” used by all living things. Pigments within these autotrophs (auto = self, troph = nourish) help to capture the energy from the Sun.

After completing this laboratory, students should be able to:

• Separate pigments by thin-layer chromatography and calculate the R_f value
• Describe a technique to determine photosynthetic rates
• Compare the photosynthetic rates of active chloroplasts and heated chloroplasts using controlled experiments
• Explain why the rate of photosynthesis varies under different environmental conditions

Concepts

• Photosynthesis
• Adsorption
• Plant pigments
• Chromatography
• Separation of mixtures

Background

I. Photosynthetic Pigments

A pigment is a molecule or compound that absorbs certain wavelengths of visible light very strongly and transmits or reflects the remaining wavelengths of visible light. Different pigments have different molecular structures, and thus reflect different specific wavelengths of visible light. The characteristic color of an organism is due to the reflection of specific wavelengths of light by pigments within the organism (see Table 1). The major pigments of photosynthetic organisms are the chlorophylls. There are two types of green chlorophyll, called chlorophyll a and chlorophyll b. Chlorophyll a and chlorophyll b are found in all plants, most algae and some bacteria. Chlorophylls c and d are not green and only occur in some algae and bacteria.

*appear grey-yellow when seen in low concentrations.

In addition to the chlorophylls, autotrophs also contain other pigments used to collect light energy (see Figure 1). These other pigments are known as accessory pigments. There are over 600 known accessory pigments. The accessory pigments may be divided into four classes: the anthocyanins, the carotenoids, the phycobilins and the xanthophylls. The phycobilins are red to blue in color and only occur in Cyanobacteria and Rhodophyta. In Table 1, the two major phycobilins are listed; they are phycocyanin and phycoerythin. The anthocyanins are blue to purple in color and occur in plants, algae, bacteria and diatoms. Table 1 lists the pigments ending in –cyanin are anthocyanins. The xanthophylls are yellow to red in color and occur in plants, algae, bacteria and diatoms. In Table 1 the pigments ending in –anthin are xanthophylls. The carotenoids are yellow, orange or red in color and occur in plants, algae, bacteriaand diatoms. The colors of the accessory pigments are masked in plants during the summer by the high concentration of chlorophyll a. In autumn, the decrease in the amount of daylight causes the plant to break down the chlorophyll, which allows the brilliant reds, oranges and yellows of the accessory pigments to become visible in the fall leaves. The differences in the molecular structure of various pigments mean that the pigments all have different polarities as well. Variations in polarity make it possible to separate plant pigments using a process known as chromatography. There are many different types of chromatography but most work on the principle of adsorbtion. An adsorbent is a solid which is capable of attracting and binding the components in a mixture (see Figure 2). In this laboratory, a thin layer of silica spread onto a thin plate of nonreactive plastic will act as the adsorbent. Many other chemicals may also be used as the adsorbent in thin-layer chromatography (TLC). Other examples of chromatography used by scientists to separate molecules based upon the concept of adsorbtion are paper chromatography and gas chromatography. In this laboratory, a suspension of plant pigments will be “spotted” onto the surface of a silica TLC plate and a solvent will then allowed to seep or flow through the silica on the surface of the TLC plate separating the different pigments. The separation occurs because one of the components in the mixture is more strongly adsorbed onto the silica than another. As a result it will spend a smaller fraction of time free in solution and will move up the TLC plate more slowly than the solvent. Components that are not strongly adsorbed onto the silica will spend a greater fraction of time free in solution and will move up the TLC plate at a faster rate. This “partitioning” of the components of a mixture between the silica and the solvent separates the components and gives rise to different color bands that become visible on the surface of the TLC plate.

How far each pigment migrates depends upon many factors, including how high the solvent is allowed to rise, the type of adsorbent, the type and concentration of the solvent, the temperature of the experiment, and the distance of the starting point from the pool of solvent. In order to compare values, scientists calculate the relative mobility of each pigment. Scientists report the experimental conditions and the relative mobility, called the retention fraction (R_f), for each pigment and compare their results with those from other scientists conducting similar experiments. R_f is defined as the fractional rise of the pigment compared to the rise of the solvent. Written as an equation this becomes: II. Photosynthesis

The second part of this laboratory activity pertains to the process of photosynthesis. Photosynthesis is a complex process in which light energy is converted to chemical energy in the form of carbohydrates and sugars. Of all of the pigments, only chlorophyll a directly captures light energy and converts it to chemical energy. Chlorophyll b and the accessory pigments assist the process by expanding the range of wavelengths absorbed (see Figure 1) and then transmitting the energy to chlorophyll a.

When light energy is absorbed by chlorophyll a, it boosts electrons within the chlorophyll molecule to a higher energy level. This extra electron energy is used to produce ATP and to reduce nicotinamide adenine dinucleotide phosphate (NADP) to NADPH. In a higher order plant, the reduction occurs within the thylakoid of the plant’s chloroplasts (see Figure 3) and is called the light reaction. A thylakoid is a saclike membrane in the chloroplast. After being produced in the light reaction, ATP and NADPH are subsequently used to incorporate carbon dioxide into glucose in a process called carbon fixation. Carbon fixation occurs in the fluid of the chloroplast that surrounds the thylakoids. This fluid is called the stroma (see Figure 3). Carbon fixation is also called the dark reaction or the light independent reaction. Chloroplasts are membrane-bound organelles are found only in eukaryotes. In Eubacteria, which are prokaryotes, photosynthesis occurs within thylakoids found within folds of the cell membrane called chromatophores. If these thylakoids contain chlorophyll the process of photosynthesis is similar to that found in plants and algae. In the prokaryote kingdom Archae, also known as Archaebacteria, some species conduct photosynthesis in a very different manner because they lack chlorophyll. These species use hydrogen sulfide or salt instead of water to capture the energy from the Sun.

In order to study photosynthesis, scientists have developed methods to observe the transfer of energy through the light reaction. One compound that allows scientists to monitor the absorbance of light energy by chloroplasts is 2,6-dichlorophenolindophenol (DCPIP). DCPIP can be used in place of the electron acceptor NADP in photosynthesis. DCPIP is useful because it is a dye that intercepts the flow of electrons in the photosynthetic electron transport chain. The dye accepts the electrons and becomes reduced, causing a visible change in color—from blue (oxidized form) to colorless (reduced form). This color change can be measured using a spectrophotometer as a reduction in absorbance or an increase in transmittance through the optical cell. In order to substitute DCPIP for NADP, chloroplasts will be extracted from spinach leaves and then incubated with DCPIP in the presence of light.

Experiment Overview

In Activity 1, pigments extracted from spinach leaves and blue-green algae (Cyanobacteria) will be separated and compared using thin layer chromatography. The retention fraction (R_f) value for each pigment will be calculated and each pigment will be identified.

In Activity 2, the rate of photosynthesis will be compared in samples with active chloroplasts exposed to light and those that are not exposed, also heated chloroplast will be compared to the unheated–active chloroplasts.

Materials

Safety Precautions

Acetone and the chromatography solvent are flammable and dangerous fire risks. They are also toxic by ingestion and inhalation. This lab should be performed only in an operating chemical fume hood or well-ventilated area. Wear chemical splash goggles, chemical-resistant gloves and a chemical-resistant apron. Wash hands thoroughly with soap and water before leaving the laboratory.

Procedure

Activity 1. Identification of Photosynthetic Pigments

1. Label two 50-mL beakers with your group name or number. Label one beaker “spinach extract” and the other “blue-green algae extract.”
2. Touching only the edges of the TLC sheet, use scissors to cut the TLC sheet into small plates approximately 6.5 cm x 1.5 cm. Be careful not to scrape any of the silica gel off the plates—this will adversely affect results. Note: Some silica gel may chip from the edges of each plate—this is not a problem.
3. Using a pencil, draw a faint line 0.5 cm from the bottom edge of each TLC plate (see Figure 4).
   {10779_Procedure_Figure_4}
4. Fill the stem of a thin-stemmed pipet with one of the extracts.
5. Carefully drip one small drop of extract onto the center of the pencil line on the TLC plate. Note: It is important to keep the spot centered and as small as possible.
6. Blow gently on the spot to help the solvent evaporate.
7. Repeat steps 5 and 6 five to ten times in the same spot to create a dark spot of extract on the TLC plate. Note: Too much extract on the initial spot will cause the resulting pigment bands to broaden and tail. Too little extract will create very faint bands that are difficult to analyze.
8. Repeat steps 4 to 7 using the second extract and the second TLC plate.
9. Use a graduated pipet to transfer 2 mL of chromatography solvent into the bottom of each 50-mL beaker.
10. Carefully place each TLC plate, sample end down, into the appropriate 50-mL beaker. Note: The upper level of the solvent must not touch the sample extract (see Figure 5). If it appears that the sample will be within the solvent, take out the sample before the TLC plate touches the solvent and remove some of the chromatography solvent from the 50-mL beaker.
    {10779_Procedure_Figure_5}
11. Carefully, place the watch glass onto the beakers. Note: Do not disturb the TLC plate.
12. When the solvent front (top of the solvent) is 0.5-1 cm from the top of the TLC plate, remove it from the 50-mL beaker. Replace the watch glass on top of the 50-mL beaker to prevent the solvent from evaporating.
13. Use a pencil to immediately mark the location of the solvent front before it evaporates.
14. Mark the center of each pigment band.
15. Measure the distance each pigment migrated from the sample spot to the center of each separated pigment band. Record the distance, in millimeters, that each pigment and the solvent front moved on the Chromatography Worksheet.
16. Calculate the R_f value for each pigment and record this value for each pigment on the Chromatography Worksheet.
17. Refer to Table 1 in the Background section to identify each pigment band. Record the identity of each band on the Chromatography Worksheet.

Activity 2. Photosynthesis

1. Turn on the spectrophotometer or colorimeter and allow to it warm up as directed by the instructor. Note: On a spectrophotometer, set the wavelength to 600 nm by adjusting the wavelength control knob.
2. Use a marker or wax pencil to label the top rim of six cuvets with zero, blank, control, active, dark or heated.
3. Using lens tissue, wipe the outside walls of each cuvet. Do not wipe off the labels. Note: Handle cuvets only near the top rim.
4. Using a square of aluminum foil, cover the outside walls and bottom of the dark cuvet. Use a second square of aluminum foil to make a loose lid for the cuvet. Note: The dark cuvet will contain the same components as the active cuvet but the aluminum foil will shield the chloroplasts from light.
5. Use the 5-mL syringe to add 5 mL of phosphate buffer to the zero cuvet and the control cuvet.
6. Rinse the syringe three times with deionized water.
7. Use the 5-mL syringe to add 5 mL of the heated chloroplast suspension to the heated cuvet.
8. Rinse the syringe three times with deionized water.
9. Use the 5-mL syringe to add 5 mL of the active chloroplast suspension to the blank cuvet, the active cuvet, and the dark cuvet.
10. Zero the spectrophotometer until the meter reads 0% transmittance.
11. Insert the zero cuvet into the sample holder and adjust the instrument to 100% transmittance. Note: The zero cuvet is actually what scientists call an instrument blank. It will be used to zero (recalibrate) the instrument before each reading. Make sure that all of the cuvets are inserted into the sample holder with the front mark on the cuvet facing the front mark on the instrument each time.
12. Using the blank cuvet, quickly complete the following steps (12 a–e)
    1. Use the 1-mL syringe to add X mL of deionized water to the blank cuvet (amount, X, determined by teacher).
    2. Immediately cover the blank cuvet with Parafilm and invert to mix.
    3. Remove the Parafilm and insert the blank cuvet into the spectrophotometer’s sample holder and read the percent transmittance. Record the percent transmittance and the time on the clock in the 0 minute box in the data table on the Photosynthesis Worksheet.
    4. Place the blank cuvet into the incubation test tube rack.
    5. Record the percent transmittance of the blank cuvet again after 5, 10 and 15 minutes have elapsed. Proceed with the rest of the lab while waiting to take the next reading. Note: Mix the cuvet's contents just prior to each reading. Use a clean piece of Parafilm each time. Remember, use the zero cuvet to check and adjust the spectrophotometer to 100% transmittance before each reading.
13. Using the control cuvet, quickly complete the following steps (13 a–e).
    1. Use the 1-mL syringe to add X mL of DCPIP to the control cuvet (amount, X, determined by teacher).
    2. Cover the control cuvet with Parafilm and invert the cuvet several times to mix the solution.
    3. Remove the Parafilm and insert the control cuvet into the spectrophotometer’s sample holder and read the percent transmittance. Record the percent transmittance and the time on the clock in the 0 minute box in the data table on the Photosynthesis Worksheet.
    4. Place the control cuvet into the test tube rack. Place the test tube rack in the incubation area as shown in Figure 6. Note: Ensure each cuvet is not in shadow.
       {10779_Procedure_Figure_6}
    5. Record the percent transmitaance of the control cuvet again after 5, 10 and 15 minutes have elapsed. Proceed with the rest of the lab while waiting to take the next reading. Note: Mix the cuvet's contents just prior to each readings. Remember, use the zero cuvet to check and adjust the spectrophotometer to 100% transmittance before each reading.
14. Using the heated cuvet, quickly complete the following steps (14 a–e).
    1. Use the 1-mL syringe to add X mL of DCPIP to the heated cuvet (amount, X, determined by teacher).
    2. Immediately cover the heated cuvet with Parafilm and invert to mix.
    3. Remove the Parafilm and insert the heated cuvet into the spectrophotometer’s sample holder and read the percent transmittance. Record the percent transmittance and the time on the clock in the 0 minute box in the data table on the Photosynthesis Worksheet.
    4. Place the heated cuvet into the incubation test tube rack.
    5. Record the percent transmittance of the heat cuvet again after 5, 10 and 15 minutes have elapsed. Proceed with the rest of the lab while waiting to take the next reading. Note: Mix the cuvet's contents just prior to each reading. Remember, use the zero cuvet to check and adjust the spectrophotometer to 100% transmittance before each reading.
15. Using the active cuvet, quickly complete the following steps (15 a–e).
    1. Use the 1-mL syringe to add X mL of DCPIP to the active cuvet (amount, X, determined by teacher).
    2. Immediately cover the active cuvet with Parafilm and invert to mix.
    3. Remove the Parafilm and insert the active cuvet into the spectrophotometer’s sample holder and read the percent transmittance. Record the percent transmittance and the time on the clock in the 0 minute box in the data table on the Photosynthesis Worksheet.
    4. Place the active cuvet into the incubation test tube rack.
    5. Record the percent transmittance of the active cuvet again after 5, 10 and 15 minutes has elapsed. Proceed with the rest of the lab while waiting to take the next reading. Note: Mix the cuvet's contents just prior to each reading. Remember, use the zero cuvet to check and adjust the spectrophotometer to 100% transmittance before each reading.
16. Using the dark cuvet, quickly complete the following steps (16 a–e).
    1. Use the 1-mL syringe to add X mL of DCPIP to the dark cuvet (amount, X, determined by teacher).
    2. Immediately cover the dark cuvet with Parafilm and invert to mix.
    3. Remove the Parafilm and remove the dark cuvet from the foil sleeve and insert it into the spectrophotometer's sample holder. Measure and record the percent transmittance and the time on the clock in the 0 minute box in the data table on the Photosynthesis Worksheet.
    4. Replace the dark cuvet into the foil sleeve, cover and place it into the incubation test tube rack.
    5. Record the percent transmittance of the active cuvet again after 5, 10 and 15 minutes has elapsed. Proceed with the rest of the lab while waiting to take the next reading. Note: Mix the cuvet’s contents just prior to each readings. Remember, use the zero cuvet to check and adjust the spectrophotometer to 100% transmittance before each reading.
17. Consult your instructor for appropriate disposal procedures.

Student Worksheet PDF

10779_Student1.pdf