CHLOROPLAST MORPHOLOGY ASSAY PROTOCOL
MATERIALS
Glutaraldehyde, 50% stock, photographic grade (Sigma Chemical Company)
EDTA – disodium salt (RPI)
Distilled deionized water
10M KOH
Sharp-point scissors
Blunt-end forceps
Sharp-point forceps
Graduated cylinders: 1000 ml
Pipettes: 5 and 10 ml
Pipette bulb
Pipette tips: 200 ul and 1000 ul capacity
Rainin Pipettors (dial-in volume)
Matrix Multichannel Pipettor
Multichannel pipettor solution reservoirs
Nutating Mixer
Screw cap bottles, 500 or 1000 ml capacity
Beaker, 1000 ml
Stirring rod
Pasteur pipettes and bulb
Tube racks
Cold pack tube racks (Scienceware), stored at 4 deg. C.
Vials with caps, 2 ml capacity, color-coded blue and green
Nonfrosted slides, with a small mark made at the top left edge of each slide
Coverslips
Bottle of dH2O with eye-dropper cap
Pencil with unused eraser at end
Fume hood
Oven, set to 55 deg. C.
Refrigerator, 4 deg. C.
Balance, (OHaus)
pH Meter (Mettler-Toledo) and pH standards (pH = 4.00, 7.00 and 10.00)
Magnetic stirring plate
Leica Microscope with polarizing lens, connected to computer/monitor with Leica Application
software
Barcode scanner
CDs (sufficient number to burn images in duplicate)
Arabidopsis leaf specimens: mature expanded and petiole from mature expanded
PROCEDURE – REAGENTS
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1. In a fume hood and using gloves, prepare 650 ml 3.5% glutaraldehyde solution from 50%
stock glutaraldehyde: 45.5 ml stock, bring to 650 ml with ddH2O, using a 1000-ml
graduated cylinder. Store the solution a in 1000 ml screw-cap bottle at 4 deg C (stable for
1 week). This amount will be sufficient for 6 flats of leaf samples (576 samples).
2. Prepare 650 ml of 0.10M EDTA (disodium salt) solution: 24.2 g EDTA to approximately 550
ml ddH2O in a 1000 ml beaker. Add a magnetic stirring rod and place on the stirring plate.
The mixture will not be dissolved and will require at least 15 minutes of stirring (without
heat) to dissolve.
3. After standardizing the pH meter, adjust the pH of the dissolved EDTA solution to 9.00 with
10M KOH solution. Add the EDTA solution to a 1000 ml graduated cylinder and bring to
650 ml with ddH2O. Store in a 1000-ml screw-cap bottle at room temperature. The 0.1M
EDTA solution is stable for at least six weeks.
PROCEDURE – LEAF SPECIMENS
1. Place 2-ml vials in pre-chilled cold racks.
2. Open the 2010 Chloroplast Morphology Database (http://genomics.msu.edu/cgibin/chloroplast/index.cgi) and choose the Harvest option. Scan in the pot barcode.
Choose the appropriate leaf tissue type (expanded leaf tip, expanded leaf petiole, or young
leaf). Submit to the database.
3. Scan the barcode of a green 2-ml vial. From the same plant, cut a mature expanded leaf
and its petiole as close to the stem as possible, holding the leaf at its side. Cut off the
petiole at the leaf base and place it in the green vial.
4. Scan the barcode of a blue 2-ml vial. Cut a small triangular piece of the mature expanded
leaf from Step #4 at its distal end and place it in the blue vial.
5. Repeat for all Arabidopsis plants.
6. Close the Chloroplast Morphology Database.
PROCEDURE – CHLOROPLAST FIXATION
1. Transfer the leaf sample tubes to a microfuge tube rack.
2. At the fume hood (wear a lab coat and gloves for the fixation steps), use a Rainin Pipette or
the Matrix 8-channel pipettor and add 900 ul 3.5% glutaraldehyde to each 2-ml vial. It is
not necessary to change pipette tips for subsequent additions. Cap each tube after adding
glutaraldehyde, and make sure the specimen is immersed in the solution. Secure tubes in
the tube rack and place rack in a horizontal position using rubber bands on the nutating
mixer rock them gently back and forth to ensure they are uniformly immersed. Incubate
tubes in the hood at room temperature for 2 hours.
3. Using a Rainin Pipette or the Matrix 8-channel pipettor set to 1000 ul, remove all
glutaraldehyde from each tube, being careful not to aspirate the leaf portion. It is not
necessary to use a separate pipette tip for each tube.
4. Using a 5 ml pipette or the Matrix 8-channel pipettor, add 1 ml 0.1M EDTA to each tube.
Recap each vial.
5. Incubate sample tubes in the oven at 55 deg. C. for 2 hours.
6. Remove the tubes from the oven and allow to equilibrate at room temperature for
approximately 10 minutes. Add tubes to microfuge tube racks and store at 4 deg. C for up
to 3 days.
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SLIDE PREPARATION AND SAMPLE SCORING – PLANT BIOLOGY BLDG – ROOM 339
1. Turn on the microscope. Log onto the computer (Log in: 2010; PW: chlorophyll). Open
the 2010 Chloroplast Morphology Database (http://genomics.msu.edu/cgibin/chloroplast/index.cgi) and choose the Analyze option. Open the Leica Application
software program. Please see the separate section for details on the operation of the
microscope and Leica application software.
2. Add a small drop of dH2O from the eye-dropper bottle to the left side of the slide (the same
side as the small mark placed beforehand). Using blunt and sharp forceps, remove a fixed
leaf or petiole sample from the vial and place it in the water drop. Cover the sample with a
coverslip. This arrangement allows each slide to accommodate three samples.
3. Tap the top of the coverslip above the specimen with the eraser-end of the pencil to crush
the specimen to a fine green haze. Use a tissue to blot off any water that has oozed out
the sides of the slide.
4. Invert the sample and view at 40x power (the mark on the slide will now be at the left side
and closest to you). Scan the slide specimen and examine the chloroplasts within the cells,
locating several to photograph. Focus the image to be photographed using the computer
monitor; a slight lag time occurs between focusing the microscope and the focused image.
Photograph two images per sample with one or two representative cells in the fields. If
abnormal chloroplasts are seen, it is best to capture one or more additional images.
5. Scan the sample tube into the Chloroplast Morphology Database. Score the chloroplasts in
each sample for:
a. Number
b. Size
c. Shape
d. Shape characteristics
In addition, any unusual findings not captured above, or other remarks for clarification
purposes should be added to the Comments section at the bottom of the scoring form.
6. Submit the results to the database.
7. Repeat steps #1-5 for each additional sample. It is easiest to work with one leaf tissue type
at a time; this allows differences in chloroplast morphology to be more easily detected.
NOTE: To minimize sample deterioration, the following method for two persons to prepare and
view the samples may be used: one person operates the microscope, views all samples, and
verbally describes the findings. The second person prepares the slides, operates the
microscope’s image capture software, and enters the results into the Chloroplast Morphology
Database. Both persons view the computer monitor to determine the best image and focus.
NORMAL CHLOROPLAST MORPHOLOGY
Expanded leaf: 80-100 chloroplasts in each cell is normal (check several planes when
viewing). Chloroplasts are small, round and grass green.
Petiole: cells are very large and rectangular with rounded corners. One petiole cell may
extend the entire length of the microscope viewing field at 40x magnification.
Chloroplasts are numerous (80-100) as in the expanded leaf, but are in several distinct
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planes. The sample must be focused through several additional planes in order to best
quantify the number of chloroplasts.
ABNORMAL CHLOROPLAST MORPHOLOGY
Cells may appear to be normal, but the chloroplasts can be very large, irregularly shaped, or
more/fewer in number. Membranes may not be clearly defined and cells may appear to have
one or two giant pale green chloroplasts within them or numerous amorphous and/or
heterogeneous-shaped chloroplasts.
LEICA MICROSCOPE – SETTINGS AND SET-UP
The microscope is positioned on three anti-vibration pads. Do NOT move the
microscope from its position on the pads or attempt to slide the entire assembly to
another place on the counter.
Tilting the microscope: the pillar holding the camera housing may be tilted if needed to
add or remove slides to the stage. Always return it to the vertical position. There is a
knob to lock the position at the back of the microscope.
Field diaphragm: used to set the condenser. Using the 10x objective, focus the light on
the same plane. Close down the condenser to center the light in the field of view. Use
the silver knob at back to keep the edges sharp and the “circle of light” centered on
screen. Then open the condenser all the way so that the field of view fills the monitor
screen.
DLF - daylight filter (on microscope pillar): Leave in at all times – should be in the “up”
position.
Condenser: polarizing lens in, analyzer tab pushed in, prism on both sides set to
number 7.
Light stop lever: can use when changing slide samples so that light does not blind you
as you change slides.
D1 = objective lens: 1, 2, 3, and 4 = 10x, 20x, 40x, and 63x (make sure the #3 prism is
set when using the 40x objective) to view chloroplasts
For light to pass to the camera vs. eyes: keep the plunger all the way out so that the
light goes to camera; specimens are still easily viewed by eye through the ocular
lenses.
Aperture diaphragm: As the aperture diaphragm is closed down, resolution is lost.
Leave it at “PH” = wide open.
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Use ethanol on lens paper to clean ocular and objective lenses. DO NOT EVER USE
KIMWIPES OR OTHER TISSUE TO CLEAN THE LENSES – THEY WILL SCRATCH
THE GLASS!!!
LEICA APPLICATION SOFTWARE PROGRAM:
SET UP
Choose the Acquire tab. Select MIC1: make sure the magnification setting matches
the microscope objective lens: 40x. Keep the Mag changer at 1x.
Camera tab: white balance on DIC. At the beginning of each session, the white
balance must be calibrated. Find an area of a sample slide having background only (no
sample, water droplets, etc.). Left click and drag the mouse to create a box in that area.
A drop-down box will appear. Choose white balance; it will be adjusted automatically.
Image section: Iris: if highlighted, leave on autofocus
Saturation: 1.45
Gamma: 0.54
Brightness: 60%
Input options: 8 bit as JPEG. Images are saved as JPEG files.
Captured frame: full frame
Captured format: full frame
Calibration Tab: DON’T CHANGE ANYTHING!
Calibration settings: bar can move for um (microns). Make sure objective and screen
objectives match (40x) so that the micron determination is accurate.
Processing Tab: Shading - none; Sharpening – low
CAPTURING AND SAVING IMAGES
1. Create a folder on the computer desktop for that session’s images.
2. Under Options>>Preferences (or press Control P), highlight the field to name the
saved image and scan in the barcode of the sample tube. Make sure the image
is saved onto the computer’s C drive and on the newly created desktop folder.
The computer will automatically add a suffix number of 1, 2, 3, etc., to the
filename of each subsequent image from the same sample so that each sample
tube need be scanned in only once.
3. Choose the Acquire Tab. Center the cell or cells of interest in the monitor’s field
of view and focus the image, using the fine focus on the microscope.
4. Press the Acquire Image. This will capture the image and save it to the folder on
the desktop as well as storing a thumbnail image at the bottom of the screen.
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5. Select the Process tab and check the appropriate boxes to add the date, time,
image name and microns labels to the captured image. Select Merge to fuse
these items onto the image.
6. Make sure the user length is selected as 20 um for the bar label. Label text
should be white, with a gray background for the text. Make sure “Autosize” is
checked.
7. Repeat steps #3-5 for each image of that sample.
Burn 3 CDs for each. Roxio: Select all of the images for the chlorophyll folder. Check
to make sure the images are satisfactory before deleting from the desktop. It is best to
wait until the week of chloroplast morphology assays are complete before deleting the
image files.
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