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PlantingScience CC BY-NC-SA 3.0 | www.plantingscience.org | Celery Challenge—Toolkit of 22 Last Updated 7/2013 Last Updated 7/2013

BACKGROUND:

The PlantingScience Celery Challenge Toolkit provides background, materials lists, detailed procedures, and safety considerations for additional experimental methods related to osmosis, transpiration, and visualizing plant cells. These tools can provide students the opportunity to ask a wider range of research questions during the Celery Challenge inquiries than would otherwise be possible. Alternatively, teachers may select one or more of these methods as the basis for classroom demonstrations of plant physiology or morphology. CONTENTS:

Page Cutting Transverse Sections of Plant Tissues for Microscopy ....................................................................... 2 Making Epidermal Peels for Microscopy ...................................................................................................... 4 Visualizing & Counting Stomata Using the Impression Method ................................................................... 6 Visualizing Plant Cells Using a Microscope ................................................................................................... 9 Determining the Electrical Conductance of a Solution ............................................................................... 12 Measuring Transpiration Using a Simple Potometer .................................................................................. 14 Quantifying Water Mass in Celery .............................................................................................................. 17 Plant Cell Staining Techniques .................................................................................................................... 19

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PlantingScience

Celery Challenge Toolkit

Image: findingthenow (Flickr)


CUTTING TRANSVERSE SECTIONS OF PLANT TISSUES FOR MICROSCOPY

Purpose: To create thin cross-‐sections of celery tissue for microscopic visualization.

How the Method Works: Transverse sections are samples of plant tissue cut perpendicular to the long axis of the organ. For example, leaf transverse sections allow the visualization of a thin slice of internal tissues from the leaf’s upper surface to its lower surface. Here, the focus is on celery petioles and the visualization of internal structures such as vascular bundles and pith. The first method, hand sectioning, is fast and simple, but requires more skill. The second method describes how to build a “poor man’s microtome” – an instrument that can be used to cut very thin sections of plant material. Sections can then be examined with a microscope. If desired, use Plant Cell Staining Techniques (p. 19) to identify cell types and components.

Technical Complexity: Simple for hand sectioning, moderate for the microtome.

Time Required: 5 minutes for hand sectioning; 10 min. for building and using the microtome.

Materials: Hand Sectioning: Poor Man’s Microtome: Celery petiole for observation Celery petiole for observation Single-‐edged razor blade Nut and bolt of compatible sizes Small dish of tap water Wax, in a small microwaveable container Toothpick Microwave Microscope slide Utility knife (Optional) Potato or carrot Microscope slide

Hand Sectioning: Hand sectioning is best used on relatively firm tissues and may be difficult if celery is limp. This method takes practice but can produce excellent results.

1. Prepare a small dish of water, then select the desired plant specimen and a new razor blade.

2. Hold the part to be sectioned firmly between your thumb and index finger (see image). Sandwich small or fragile parts between pieces of potato or carrot.

3. Taking care to avoid cutting yourself, make a preliminary cut in the plane of tissue you eventually want to examine.

Make a Hand Section

Make an Epidermal Peel


• Slice towards yourself with the razor, using as much of the blade surface as possible. • This cut will expose fresh tissue; you can discard the older piece it removes.

4. Using the same cutting technique, quickly make a series of sections, each as thin as possible. 5. Place each section in your dish of water as it is made.

• Avoid tearing the section as you move it. An ideal section includes the entire cross-‐section of tissue.

• Keep the pieces, however! These are often the thinnest. 6. Lay out a glass slide and place a drop of water onto it using a disposable pipette. 7. Transfer the thinnest section to the slide, using the flat side of a toothpick if needed. 8. (Optional) If you want to stain the cells, use a procedure from Plant Cell Staining Techniques.

Poor Man’s Microtome: Although microtomes can be purchased, they are usually quite expensive. This method can be used much more cheaply and will give better results than can be achieved by hand sectioning alone.

Steps in Using a “Poor Man’s Microtome,” as Described in the Instructions Below

1. Cut a small, cubical piece of plant material, such as from a stem. 2. Place the plant piece into a nut that is barely fastened to a bolt. 3. Melt a small amount of wax in a microwave for about 2 min, then pour the melted wax onto the

plant material, filling the space around it in the nut. 4. After the nut is filled completely, let it sit for about 2 min while the wax cools and hardens. 5. With the bolt on its side, cut straight down the nut with a utility knife to make a thin section.

• In all steps using the utility knife, take proper care to avoid cutting yourself. • This first segment of plant tissue plus wax can be discarded.

6. Screw the nut slowly onto the bolt to raise a thin section of fresh wax and tissue, then cut again. • Repeat until you have enough thin sections to work with.

7. Discard the wax from the section(s), and place the best section on a slide. 8. Place one drop of water onto the plant section or (optionally) carry out a staining procedure.

Step 1 Step 2 Step 3

Step 5 Step 7 Step 8


MAKING EPIDERMAL PEELS FOR MICROSCOPY Purpose: To remove samples of the outer cellular layer of plant tissue for microscopic visualization.

How the Method Works: Epidermal cells make up the “skin” of a plant. The cells form a distinct tissue layer that restricts water movement out of the plant; this layer also contains specialized cells called stomata that regulate gas exchange and transpiration.

Quantifying and measuring epidermal and stomatal cells is only possible with a microscope, and this can provide information about the plant’s ability to move water. Two methods for collecting epidermal samples for microscopy are described here. Epidermal peels are fast and make excellent samples for quantification, but they require some skill. Epidermal scrapes are much easier, but they collect only a few cells at a time. If desired, use Plant Cell Staining Techniques (p. 19) to identify cell components.

Technical Complexity: Simple.

Time Required: 5 minutes.

Materials: • Celery petiole or leaf • Single-‐edged razor blade or scalpel • Toothpick • Microscope slide

Epidermal Peels: Single layers of epidermal cells can be obtained using this procedure. It will likely be easier to remove the epidermis from the smooth, concave side of the celery than from the ridged, convex side.

1. Break the petiole in half so that only the epidermis connects the two pieces. o Cutting partway through the opposite side from the one you are sampling will help, as

shown in the figure above. Take care to avoid cutting yourself. 2. Gently pull one half of the petiole down the side of the other so that the epidermis is pulled

away from the tissue of the latter. 3. Lay the exposed epidermis on a drop of water and cut away the remaining tissue. 4. Use the flat side of a toothpick to transfer the exposed epidermis to a microscope slide. 5. (Optional) If you want to stain the cells, use a procedure from Plant Cell Staining Techniques. 6. (Optional) Assess the density and condition of stomata and epidermal cells with the last section

of Visualizing & Counting Stomata Using the Impression Method, and quantify the sizes of cells and stomatal pores using Steps 5-‐7 of Visualizing Plant Cells Using a Microscope.

Make a Hand Section


Make a Hand Section


Make a Hand Section



Epidermal Scrapes: Individual cells, or small groups of cells, can be obtained by scraping the celery surface. If you have difficulty getting epidermal peels from the ridged side of a celery petiole, this alternative may be helpful. Unfortunately, you will not be able to measure cell density with this approach.

1. Taking care to avoid cutting yourself, use a scalpel, razor, or wooden toothpick to scrape the epidermal tissue you wish to examine.

2. Rinse the scraper in a drop of water on a microscope slide. 3. (Optional) If you want to stain the cells, use a procedure from Plant Cell Staining Techniques. 4. (Optional) You can quantify the sizes of cells and stomatal pores using Steps 5-‐7 of Visualizing

Plant Cells Using a Microscope.


VISUALIZING & COUNTING STOMATA USING THE IMPRESSION METHOD

Purpose: To determine stomatal density and examine the state of stomata in leaves by taking impressions.

How the Method Works: Painting nail polish onto the surface of a plant tissue creates an impression of its outer cellular structure. The impression is peeled from the tissue, then examined under a microscope to view the stomata and count how many are present per unit area. You may also determine areas of guard cells, stomatal pores, and epidermal cells, or cell volumes before and after an experiment using Steps 5 through 7 from Visualizing Plant Cells Using a Microscope.

Technical Complexity: Moderate. It can take some practice to learn how to bring specimens into focus without damaging the lenses.

Time Required: 60 minutes from set-‐up to completion.

Materials: • Celery samples (1-‐2 per treatment) • Clear nail polish • Compound light microscope • Blank microscope slides • Dissecting probe or other pointed instrument • Forceps • Distilled water and eyedropper • Permanent marker • (Optional) Prepared slide containing a known specimen • (Optional) Digital camera

Lab Safety: Broken coverslips and broken slides are very sharp. Dispose of these materials in the glass disposal, NOT the regular trash can. Wearing well-‐fitting lab gloves is recommended to help prevent accidental cuts.

Prepare the Samples: 1. In your notebook, describe the celery tissues you will sample.

a. Indicate for each sample whether the impression is being made before or after experimental treatment, and what that treatment is.


b. Identify whether you will be sampling a leaflet or petiole. Will leaflets be sampled on the underside, top side, or both? Will petioles be sampled on the concave side, convex side, or both?

c. Make digital photographs or sketches, using labels or file names to keep track of all samples. 2. Prepare epidermal impressions from each sample.

a. Paint a 1 cm x 1 cm square (1 cm2) of clear nail polish onto the tissue surface. Make sure there are no gaps in the layer of nail polish. Several coats are okay, since you don’t want the nail polish to tear as you peel it off.

b. Allow the nail polish to dry thoroughly.

Get the Microscope Ready: Set up the microscope while waiting for the nail polish to dry. Ask your teacher for assistance. You may wish to learn more about safely using the microscope or refresh your memory by reading the Microscopy Manual.

1. Turn the microscope light on. 2. Move the stage far from the objective using the rough focus

knob. 3. Push the lowest power objective into place. 4. (Optional) Practice with a slide containing a known specimen to

help you familiarize yourself with the instrument.

Set Up Slides of the Impressions: 1. Gather one microscope slide for each impression. 2. Lift off each nail polish square – the tissue impression – as one

piece. a. Use a dissecting probe to gently tease the edge of the nail

polish up, lifting or peeling it away from the celery tissue until about halfway lifted.

b. Use forceps to finish peeling away the square. c. Remember which side of the peel was facing the tissue.

3. Place a drop of distilled water onto a microscope slide. 4. Put the impression onto the surface of the water with the side that was touching the tissue

facing up, away from the water. 5. Gently smooth out the impression using the dissecting probe so that it lays flat against the slide. 6. With a permanent marker, label the slide to indicate basic information about what tissue the

impression came from. o For example, you might record a sample number on both the slide and the description of

the sample in your lab notebook. 7. Repeat steps 2-‐6 with the remaining impressions.

Visualize the Impressions:

The large donut shape above is an impression of the two cells that form a stomate. Stomata are openings through which gases enter and leave the leaf. The cells around the pore, called guard cells, open and close to make it bigger or smaller.


Examine each impression under the microscope and collect data on the stomata. Remember that you are looking at a clear impression of the tissue surface, not the tissue itself!

1. Take notes and make sketches or digital photos of what you see. 2. Find the impressions of the stomata.

a. Are they open, closed, in between, or a mix? Take notes. b. Make a sketch or digital photo of an average stomate in the impression.

3. Count the number of stomata in the entire impression and record the number. o Have one other teammate count all stomata in the same slide to confirm the number. o Stomatal density is the number of stomates per cm2, so counting over the 1 cm2 impression

gives a direct estimate of this value. o Stomatal number over the whole tissue is stomatal density times tissue surface area in cm2.


VISUALIZING PLANT CELLS USING A MICROSCOPE Purpose: To use microscopy to quantify the size and density of different plant cells.

How the Method Works: Wet mount slides of celery cells are viewed at 400X magnification, with subsamples measured to calculate size, volume, and density of different cell types.

Technical Complexity: Moderate. You may need to make several samples to get a thin enough layer to see individual cells clearly. It can take some practice to learn how to bring specimens into focus without damaging the lenses and to accurately measure cells using optical tools.

Time Required: 30 minutes.

Materials: • Compound light microscope with 40X objective and at least one lower-‐powered objective • Lens paper • Samples of treated celery tissue • Scalpel • Microscope slides and coverslips • Eyedropper or small pipette • Water • Forceps • Ocular micrometer • Stage micrometer or ruler • (Optional) Two dissecting needles • (Optional) Digital camera • (Optional) Polarizing filters

Lab Safety: Scalpels, broken coverslips, and broken slides are very sharp. Dispose of these materials in the “sharps” container or in the glass disposal, NOT the regular trash can. Wearing well-‐fitting lab gloves is recommended to help prevent accidental cuts.

Procedure: 1. Before you begin, carefully clean the objectives and eyepieces of your microscope with lens paper.

2. If you have not already done so using one of the previous three methods, prepare your specimen(s): a. Select a tissue of interest and use a scalpel to slice a very thin fragment from it.

PlantingScience CC BY-NC-SA 3.0 | www.plantingscience.org | Celery Challenge—Toolkit of 22 Last Updated 7/2013

b. Place a drop of water into the center of a microscope slide using an eyedropper or pipette, then transfer the fragment to the water on the slide.

c. Gently tease apart the tissue in the water droplet using the scalpel or two dissecting needles.

d. Place one end of a glass coverslip to the right or left of the specimen so that the rest of the slip is held at a 45o angle over the specimen.

e. Slowly lower the coverslip with a dissecting needle or forceps, removing any air bubbles. The coverslip flattens the sample, keeps it from drying out, and protects the

objective lenses. f. Press down very, very lightly on the cover slip with the dissecting probe or forceps.

This spreads and flattens the tissue, so that you can better see one layer of cells.

3. Set up and calibrate an ocular micrometer to take accurate measurements with the microscope: a. If you have not used a microscope recently, make sure you ask your teacher and read up on

using them. b. Insert the micrometer into the microscope’s ocular lens.

You may need to unscrew the top element of the ocular lens to do this. If no top element exists, you will need to hold the micrometer in place with a split ring.

c. Place a stage micrometer or plastic ruler onto the stage for calibration. d. Using the lowest power objective, bring the scale on the stage micrometer or ruler into focus. e. Line up one end of the stage micrometer or ruler with one end of the ocular micrometer scale. f. Read across the scales until you find two lines that are exactly superimposed. g. Count the number of scale units on the ocular micrometer needed to reach this point. Record

this and the corresponding measurement in mm from the stage micrometer or ruler. h. Divide the actual measurement by the number of ocular micrometer scale units to determine

the distance measured by one ocular micrometer scale unit. i. Repeat this procedure for the other objectives you plan to use.

4. Observe your specimen(s) under the microscope: a. Begin by using the lowest-‐power objective. b. Locate the part of the slide that has seems to have the fewest

layers of cells. c. Use the focus to help distinguish among individual cells.

Where are the cell walls? What cell types are present? What are their distinguishing features? Make a sketch of your observations, noting the total magnification in each drawing. Record notes about the sample. For example, to what treatment was it subjected? Alternatively, use a digital camera to take microscope photos, writing the total

magnification, sample information, and photo file name in your lab notebook. You may wish to use a polarizing filter to distinguish between the evenly thick secondary

cell walls of sclerenchyma and the thinner primary cell walls of other cell types.


d. Use the 40X objective to look at the sample. Again, make a sketch or digital photo of your observations. Try to answer the following questions in your lab notebook:

Where are the cell walls? What cell types are present and what are their distinguishing features? What details do I see at this scale that I did not notice at lower magnification?

5. Determine cell area for 3-‐5 cells of a given type: a. Use the objective lens that allows you to take precise measurements of a given cell type. b. Measure width and length of cells that are typical of what you see in the sample using the scale

units of the ocular micrometer. You may need to reposition the sample to do this. Be sure you measure the same cell

for both length and width! c. Record the number of scale units in your lab notebook. d. Convert the scale units to measurements in mm using the results from your calibration in Step 3. e. Calculate the average width and average length of these cells. f. Calculate average cell area as average cell width x average cell length. g. Direct measurements in different cell types will be difficult to compare, so you may calculate %

change in average cell area before and after an experiment as 100% x (!"!#!$% !"#!!!"#$% !"#!)!"#$% !"#!

.

6. Determine cell volume for the same cells you used for Step 5: a. Assume that the cells are basically cubiodal. Given the average area of this cell type, multiply it

by average length to give one “endpoint” estimate of the cell volume. b. Multiply the average area of the cell type by average width to give the other “endpoint”

estimate of cell volume. c. The true average cell volume is likely to fall between these two values. d. Direct measurements across cell types will be difficult to compare, so you may also calculate %

change in average cell volume as 100% x (!"!#!$% !"#$%&!!"#$% !"#$%&)!"#$% !"#$%&

.

7. (Optional) Using an epidermal sample, determine average stomatal pore size from 3-‐5 stomates: a. Use the objective that allows you to take precise measurements of a stomatal pore. b. Adjust the stage to position a stomate so you can measure its pore length and width using the

ocular micrometer. Record the length and width in your notebook. c. The approximate stomatal pore area can be described as pore length x pore width. d. Repeat for additional stomata to determine the average stomatal pore area. e. The proportion of total tissue area accounted for by stomatal pore area may also give you an

idea of how much of the tissue surface allows gas exchange: 𝑎𝑣𝑒𝑟𝑎𝑔𝑒 𝑠𝑡𝑜𝑚𝑎𝑡𝑎𝑙 𝑝𝑜𝑟𝑒 𝑎𝑟𝑒𝑎 x 𝑎𝑣𝑒𝑟𝑎𝑔𝑒 𝑠𝑡𝑜𝑚𝑎𝑡𝑎𝑙 𝑑𝑒𝑛𝑠𝑖𝑡𝑦

You must also determine stomatal density in your specimen using the last section of Visualizing & Counting Stomata Using the Impression Method to make this estimate.

You need not calculate the total surface area of the tissue from which you took the specimen, because stomatal density is already adjusted per unit area.


DETERMINING THE ELECTRICAL CONDUCTANCE OF A SOLUTION

Purpose: Use conductivity to quantify the amount of ions in the salt solution you have used or will use to soak celery samples.

How the Method Works: Electrical conductance, or conductivity, is the amount of electricity that can be conducted over a given

distance in an aqueous solution. This measure is often used to represent the salinity, or saltiness, of a liquid solution. Handheld meters can be used to accurately measure a solution’s conductivity, and therefore salinity, even after some evaporation has occurred.


Time Required: 10 minutes per reading.

Materials: • Aqueous solution containing one or more types of ionic solute • Handheld digital conductivity meter • One or more conductivity standards • Distilled water • Small containers for standards and sample

Calibrating the Conductivity Meter Conductivity meters will differ based on the manufacturer. Be sure to read the instructions that come with the instrument, and use those if they conflict with any of the steps below.

1. Turn on the conductivity meter and allow it warm up for at least five minutes. 2. If there is one, adjust the temperature knob to match the solution temperature. 3. Calibrate the meter against one or more standards having known conductivity.

a. Pour just enough of a standard into its own container so that it will immerse the meter’s electrode.

b. Insert the electrode into the standard and allow the meter to reach a steady reading. c. If the meter does not read the same as the salinity of the standard you used, adjust the

meter to set it to the proper salinity. Follow the manufacturer’s instructions as needed. d. Rinse the meter with distilled water, collecting the rinse water in another container. e. Hold the tip of the electrode against the outside of the container to allow any large drops of

water to flow off the electrode. f. If desired, repeat steps a-‐e with additional standards to calibrate the meter over a larger

range of conductivity values.


Measuring Sample Conductivity 1. Rinse the electrode with distilled water as described previously. 2. Insert the electrode into the unknown solution. 3. Allow the reading to stabilize before recording the value. 4. When you take the electrode out of the solution, rinse it again with

distilled water. 5. Cover the tip or place the electrode back into its proper storage

conditions.


MEASURING TRANSPIRATION USING A SIMPLE POTOMETER Purpose: Quantify the amount of water a celery leaf transpires by monitoring its water uptake.

How the Method Works: Transpiration is the movement of water through a plant. The loss of water through stomatal pores in the shoots is also considered part of transpiration, as this evaporative process, along with osmosis, drives the upward movement of water in the plant. While it is difficult to accurately measure the loss of water from a plant’s shoots,

measuring the resulting water uptake through the roots, stem, or thick petioles is simple. Here transpiration is measured in a celery leaf by placing the cut petiole into one end of a piece of tubing and sealing it. Filling the tube with water allows the measurement of water consumed by the leaf over time as the change in water volume inside the tubing.

Technical Complexity: Medium, since the process of making and keeping a seal in the tube can be difficult.

Time Required: 20-‐30 minutes per reading.

Materials: • Whole leaves or bunch of celery • Paring knife, scalpel, or single-‐edged razor blade • At least 40 cm of clear, flexible plastic tubing with a diameter large enough to fit a celery petiole • Rubber stoppers of various sizes • Parafilm or other hydrophobic sealing material • Shallow basin • Tap water • Paper towels • Permanent marker • Clock or timer • Ruler

Assembling the Potometer: Proper sealing of the potometer is the most important factor in making accurate measurements here.

1. Fill a shallow basin with tap water. 2. Submerge a length of clear tubing entirely in the basin.

o Gently move the basin or tubing until all air bubbles have escaped the tubing. 3. Remove one leaf of celery from the bunch and, taking care not to cut yourself, use the paring

knife or other sharp tool to cut the end of the petiole.

Image: Matthew McVickar (Flickr)


4. Immediately immerse the cut end of the petiole in the basin of water, and work carefully to fit the cut end into one end of the tubing. o Exposing the cut end to air too long will allow air to be pulled

into the bottom of the petiole, slowing the normal rate of transpiration.

o It may be helpful to have several sizes of tubing available or to cut the petiole further towards the leaflets in order to fit the petiole inside.

5. Close the gap between the concave side of the petiole and the rest of the tubing by fitting a rubber stopper into the same end of the tubing. o You may need to try more than one stopper size to find one that fits. o Avoid crushing the end of the petiole to prevent damage to the vascular tissues.

6. Carefully seal the joints between the tubing, celery, and stopper with hydrophobic film. a. Cut a piece of film 2-‐5 cm long from the roll. b. Place one edge of the film against the stopper and tubing. c. Pressing down on the edge of the film, use your other hand to stretch the film and pull it

around both the tubing and the stopper, and then both the tubing and the petiole. d. Wrap the film around the celery, stopper, and tubing several times until it is completely

used up. e. You may want to wrap each cycle further upwards, so that you also close the gap between

the stopper and celery.

Measuring Transpiration: This step will require two people. One person will hold the free end of the tubing and make measurements, while the other person will hold the celery leaf upright.

1. Lift the free end of the tubing above the surface of the water. 2. Pour out a small amount of water from this end, so that the meniscus is clearly visible inside. 3. The second person should lift the sealed end of the tubing above the surface of the water,

checking to make sure no leaks or air bubbles show up inside. o The tubing should now form a “U” shape, with one person holding each end. o If any leaks or air bubbles form, the seal is leaky. Go back to Steps 4, 5, and 6 in the previous

section to make a better seal. 4. If the seal is good, wipe the free end of the tubing with paper towel to dry it, then use a

permanent marker to mark the location of the meniscus. 5. Record the time at which the meniscus was marked. 6. Allow 15 to 20 min to allow for measureable levels of plant transpiration.

o If you like, you may apply environmental treatments at this time, such as different lighting, temperature, or air flow conditions.

7. Using a permanent marker, mark the meniscus position at the end of the monitoring period. o If the mark is not easily distinguished from the initial one, you may extend the monitoring

period to make the difference easier to see. 8. You may now lay down the tubing, making sure to let the water flow into the basin.


Calculating Water Consumption Rate: 1. Using a ruler, measure the length (L) in cm that the meniscus moved in the tubing during the

monitoring period. 2. Measure the internal diameter (d) in cm of the tubing at its free end.

3. Calculate the volume of the water column that was consumed as: 𝜋(!!)!𝐿 .

o The units of the calculation will be in cm3, which is equivalent to mL. 4. Divide the volume of water used by the number of minutes you monitored to

determine the water consumption rate in mL/min.

d

L


QUANTIFYING WATER MASS IN CELERY

Purpose: To determine the mass or changes in mass of water in celery petioles for osmosis experiments.

How the Method Works: There are two ways to measure plant water mass: mass change or direct quantification. In the mass change approach, the whole sample is weighed before and after any treatment short enough to allow measureable changes water mass but

not plant growth. For the direct quantification approach, samples are weighed at the final experiment endpoint, then dried and weighed again. The difference in mass between the two measurements is the water mass. Samples that are weighed for dry mass cannot be used for further studies, so you need to decide which method(s) is/are suited to your research question.


Time Required: About 5 min per plant. For dry weight measurements, an overnight drying time is required.

Materials: • Digital balance with milligram (0.001 g) precision • Paper towels • Drying oven • (Optional) Paper lunch bags • (Optional) Ziploc sandwich or gallon-‐sized bags

Measuring Change in Water Mass This approach assumes that the only variable influencing the mass of the celery is the amount of water it contains. If this assumption is false, it may be better to use direct quantification.

1. Just before starting the experiment, weigh the celery sample on a digital balance and record the mass in your lab notebook.

2. Carry out the experiment according to plan. 3. After the treatment is complete, remove the celery sample from the soaking solution. 4. Blot the sample gently with a paper towel to remove any free surface moisture. 5. Weigh the sample immediately on the digital balance.

o Plants have a high water composition, so waiting to weigh them may lead to some drying and produce inaccurate data.

6. Change in water mass = Final fresh mass – Initial fresh mass.

7. Percent change in water mass = 100% x (!"#$% !"#$% !"##!!"!#!$% !"#$% !"##)!"!#!$% !"#$% !"##

.


Direct Quantification of Water Mass This method directly measures the mass of water in the celery. However, directly measuring the same sample both before and after treatment is not possible, because the method uses destructive sampling.

1. Carry out the experiment according to plan. 2. After the treatment is complete, remove the celery sample from the soaking solution. 3. Blot the sample gently with a paper towel to remove any free surface moisture. 4. Weigh the sample immediately on a digital balance. 5. Dry all samples overnight in an oven set to low heat (60oC).

o If you have many celery stalks to dry, it may be helpful to place them individually in paper lunch bags.

o Use labels or make a chart of the drying positions to keep track of their respective treatments.

6. Let the celery cool in a dry environment. o In a humid environment, the plant tissue may take up water. o A sealable plastic bag will keep moisture out if you live in a humid

climate. 8. Once the samples have cooled, weigh them on the balance.

o Plants contain mostly water, so the dried samples will weigh much less than before. 9. The water content of a plant sample can be calculated as:

o Water mass = Fresh mass – Dry mass

o Percent water mass = 100% x (!"#$% !"##!!"# !"##)!"#$% !"##

.


PLANT CELL STAINING TECHNIQUES

Purpose: To identify different cellular structures, and thereby different cell types, using biological stains.

How the Method Works: A variety of cell stains are available to highlight different cellular structures, such as the primary cell wall, secondary cell wall, starches, and lipids. Here we describe the use of four such stains that are most likely to help distinguish among different types of celery cells.


Time Required: About 10-‐20 min to make one stain stock; 5-‐10 min to prepare one specimen.

Materials: • Microscope slide(s) containing specimen(s) • Digital balance • Spoon or scoopula • Weighing boat or weighing paper • Graduated cylinder • Water • Light-‐blocking container for storing prepared stain (e.g., foil-‐wrapped or brown glass) • Eye dropper • Dust-‐free tissues, such as Kimwipes • Timer • (Optional) Cover slips • (Optional) Forceps • (Optional) Glass Petri dish • Materials for one or more of the methods below:

Method A Method B Method C Method D • Toluidine blue O powder

• Benzoic acid • Sodium benzoate • pH meter or test strips

• Phloroglucinol powder

• Ethanol • Concentrated hydrochloric acid

• Fume hood

• Sudan IV powder • Propylene or ethylene glycol

• Hot plate • Thermometer • Whatman No. 2 filter papers

• Funnel

• Potassium iodide • Iodine • Fume hood


Method A: Staining the Primary Cell Wall This method stains cellulose in the primary cell wall. Based on the thickness and evenness of the cell wall, you can distinguish between parenchyma and collenchyma cells.

1. Prepare 0.1% toluidine blue stain: a. Make 0.1 M benzoate buffer:

i. Weigh out 0.125 g benzoic acid and 0.145 g sodium benzoate using a digital balance. ii. Dissolve both in a storage container by adding water, bringing the volume to 100 mL. iii. Test the buffer pH and adjust to 4.4 as needed by adding single drops of 1 M

hydrochloric acid or 1 M sodium hydroxide. b. Weigh out 0.1 g of toluidine blue O powder on the balance. c. Add the toluidine blue O powder to the storage container, then stir or swirl to dissolve

completely, producing 0.1% toluidine blue stain. 2. Stain the specimen:

a. Using an eyedropper, transfer one or more drops of 0.1% toluidine blue stain onto a specimen on a microscope slide you have previously prepared.

b. Allow the stain to incubate for one minute. c. Absorb as much stain as possible from the slide using a dust-‐free tissue. d. Wash the specimen by flooding it with water using the eyedropper. e. Dry the specimen using a new tissue or the eyedropper. f. If needed, repeat steps d-‐e to remove excess stain. g. Add one drop of water onto the specimen to keep it hydrated. h. Touch a cover slip to one side of the water drop using your fingers or a forceps. i. Gently lower the cover slip over the specimen,

avoiding any air bubbles. 3. Examine the specimen using a microscope.

o Cellulose and pectin will be stained reddish purple. o Recall that collenchyma will have uneven, thick cell

walls. o Parenchyma will have thinner cell walls; parenchyma

with chloroplasts are chlorenchyma. o Lignin and other phenolic compounds will range in

color from blue to green.

Method B: Staining the Secondary Cell Wall This method stains lignin in the secondary cell wall. Based on the presence or absence of lignin, you can distinguish between sclerenchyma and other cell types.

1. Prepare phloroglucinol-‐HCl stain: a. Prepare 80 mL of 20% ethanol in a storage container by adding 17 mL 95% ethanol to 63 mL

water. b. Weigh out 2.0 g phloroglucinol powder using a digital balance. c. Dissolve the phloroglucinol in the 20% ethanol solution.

Toluidine blue stained creeping-‐oxeye stem

Image: 石川 Shihchuan (Flickr)


d. In a fume hood, and while wearing gloves, add 20 mL of concentrated hydrochloric acid (12 M) to the solution.

e. Gently swirl or stir the solution to mix. 2. While still wearing lab gloves, stain the specimen:

a. Use an eyedropper to add one or two drops of phloroglucinol-‐HCl stain to the specimen. Alternatively, you may pour some stain in a glass Petri dish and move the specimen to the dish for incubation.

b. Incubate for at least two minutes. c. Absorb as much stain as possible from the slide using a dust-‐free tissue.

Alternatively, transfer the specimen from the dish to the slide using a toothpick or forceps.

d. Add one drop of water onto the specimen to keep it hydrated. e. Touch a cover slip to one side of the water drop using your fingers or a forceps. f. Gently lower the cover slip over the specimen, avoiding any air bubbles.

3. Immediately examine the specimen using a microscope. o The color will fade over several minutes. o Lignin-‐containing cells, i.e., sclerenchyma, will be

stained red. o Xylem cells will form tube shapes with pointed

ends or circles, depending on the angle at which the specimen was cut.

o Sclereids will not be elongated and may instead form branches or star shapes.

Method C: Staining Lipids This method stains the waxes, fats, and oils within a cell or tissue and will allow you to more easily identify the waxy cuticle in epidermal cells. It may also stain any suberin present, which is a compound found in a structure outside the vascular bundles called the Casparian strip.

1. Prepare the Sudan IV stain: a. Measure out 100 mL of propylene glycol or ethylene glycol in a graduated cylinder and

transfer to a heatable container. b. Weigh out 0.7 g of Sudan IV powder and transfer to the container. c. While stirring, heat the solution to 100oC on a hot plate and incubate for 5-‐10 minutes. d. With a Whatman No. 2 filter paper and funnel, filter the hot solution into another container. e. Allow the solution to cool, then repeat the filtering step, transferring the solution into a

storage container. 2. Stain the specimen:

a. Prepare a solution of 85% propylene glycol or ethylene glycol in water. b. Transfer specimen(s) directly into a small container of the Sudan IV stain. c. Incubate specimen(s) in the stain for 5 min. d. Move the specimen(s) to the 85% propylene or ethylene glycol solution, gently swirling the

container to help wash away extra stain.

Stained lignin (red) in poplar branch Image: tpuukko (Flickr)


e. After about 30 sec of washing, gently rinse the specimen(s) in water. f. Transfer the specimen(s) to the microscope slide(s). g. Add a cover slip to the specimen(s) as previously described.

3. Examine the specimen using a microscope. o Any waxes, fats, or oils will be stained red. o Epidermal cells will be distinguishable by the

presence of their outer cuticle. o You may also see a red band around the outside of

some vascular tissues. This is the Casparian strip, a hydrophobic band of suberin that prevents loss of water out of the xylem.

Method D: Staining Starches This approach stains cellular starches. You may be able to distinguish among cells used for food storage (e.g., parenchyma) and other cell types (e.g., vascular cells) using this approach.

1. Prepare the IKI stain before you begin to prepare your specimens, since iodine takes a while to dissolve. a. Using a graduated cylinder, transfer 100 mL of water to the stain storage container. b. Weigh out 2.0 g of potassium iodide on a digital balance and transfer it to the storage

container. c. In a fume hood, weigh out 0.2 g of iodine and transfer it to the storage container. d. Tightly cap the bottle as the iodine dissolves to avoid its sublimation and loss from the stain.

2. Stain the specimen: a. Make sure that both the potassium iodide and iodine have completely dissolved in the stain. b. Using an eyedropper, transfer a drop of the IKI stain onto the specimen. c. Incubate for 5 minutes. d. Add a cover slip to the specimen as previously

described. 3. Examine the specimen using a microscope.

o Starches will range in color from bluish black to reddish purple.

o Longer starch polymers will tend to have a blacker color, while shorter, newly synthesized starch polymers will tend to be more reddish.

o Nuclei and cell walls will often stain reddish brown.

Staining methods based on: Yeung, E.C. 1998. A Beginner’s Guide to the Study of Plant Structure. In S.J. Karcher, Ed. Tested Studies for Laboratory Teaching, Vol. 19: Proceedings of the 19th Workshop/Conference of the Association for Biology Laboratory Education (ABLE), pp. 125-‐142.

IKI stained leek epidermis Image: Yersinia pestis (Flickr)

Casparian strip (arrow) in maize root Image: BlueRidgeKitties (Flickr)

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