Main Page/BIOL 4160/Visualizing the Photosynthetic Apparatus

From Biology Wiki
Jump to: navigation, search

Contents

Lab 02 Visualizing the Photosynthetic Apparatus

Introduction

So far, you have had an opportunity to explore the nature of light and its fates in intact photosynthetic organs (leaves). In eukaryotic cells, photosynthesis occurs in chloroplasts. Chloroplasts are a unique organelle, containing their own DNA and the full suite of biochemical reactions required to convert light energy to carbohydrate.

In this lab, you will have the opportunity to visualize the photosynthetic apparatus and become familiar with the nature of the chloroplast organelle in a variety of photosynthetic organisms. Part of the lab objective is to compare the structure of chloroplasts in situ with their structure in vitro. Most biochemical experiments on chloroplasts require that the chloroplasts be isolated (in vitro). Thus, you will be required to isolate chloroplasts from a photosynthetic tissue (spinach leaves and algal cells) and compare these in vitro chloroplasts to in vivo chloroplasts in leaves and algal organisms cultured in the Biology Department for teaching and research.

To visualize the chloroplasts, you will take advantage of a fourth fate for a light photon interacting with the photosynthetic apparatus: fluorescence.

Microscopy

With the microscope, it will be possible to identify and measure the size of chloroplasts. You will be using bright field, phase contrast, and fluorescent microscopy and should diagram (with scales) and comment on the structures you observe with the three microscopic techniques.

It will be necessary to calibrate the microscope using a micrometer slide. Kohler illumination is crucial for optimal viewing using bright field and phase contrast. You will have had experience doing this in previous biology labs, but can request assistance from the lab demonstrator as required.

Chloroplast Isolation

Practically all biochemical assays of photosynthesis use isolated chloroplasts, in part to avoid 'contamination' with other cell organelles, especially mitochondria. Historically, careful isolation of 'pure' chloroplasts was necessary for biochemists to accurately characterize what chloroplasts do. To isolate the chloroplasts, it is necessary to break apart the tissue to the extent that individual chloroplasts are released from the cell. A typical technique to disrupt cells is a blender. It is highly effective, but has to be used with care since excessive homogenization will damage the chloroplasts (something you could assess using microscopy). An alternative is a mortar and pestle, a gentler technique that minimizes damage --we will use a mortar and pestle in this (and future) experiments. The chloroplast in situ exists in an osmotically and pH balanced environment. Thus, the tissue should be disrupted in a solution that mimics the osmolarity (about 600 mosmol/kg, sucrose is often used) and pH (about 7.2, tricine is a common buffer) of the cytoplasm. A common media used for chloroplast isolation is STK (sucrose/tricine/KCl).

STK (pre-chilled on ice):

  • Sucrose (FW 342.3) 0.6 M
  • Tricine (FW 179.2) 0.05 M
  • KCl (FW 74.55) 0.02 M)
Tricine buffer has a pK of 8.15. The pH of the media is adjusted to 7.5 with KOH/HCl, as required. The KCl is provided to mimic (not very accurately, but simply) the ionic environment of the cytoplasm.

Other materials: Spinach, Mortar and pestle (pre-chilled on ice), Algal cultures, Glass homogenizer (pre-chilled on ice), Cheesecloth (to filter the homogenates), Centrifuge and centrifuge tubes (to pellet the chloroplasts), and Miscellany (beakers, No. 1 filter paper etc.)

Pre-chilling is important because, once isolated from the tissue, the chloroplasts will be subjected to proteolytic and lipolytic activity, resulting in a progressive deterioration of the organelles. Chilling inhibits the deterioration.

Chloroplast Preparation from Spinach

Please note that this is the longest procedure you will have to undertake in this laboratory. It is also the most important, in the sense that you will be isolating chloroplasts from spinach in a number of labs. This is a case of engaging in real scientific practice: learning a procedure and repeating it until (in the words of previous students in this course) you can "isolate chloroplasts with your eyes closed" (but please don't try! Keeping your eyes open is very helpful!).

When you harvest the spinach leaves, try to minimize wilting, because wilting has an adverse effect on the yield of healthy chloroplasts. Wash the spinach leaves thoroughly under tap water, and remove midribs etc. with scissors (or a razor). Be sure to set aside leaves (still on the spinach plant) for subsequent viewing under the microscope. Cut the leaves into squares of about 2 cm. Place about 10 grams in the mortar, and cover with 50–100 ml of STK. Homogenize by crushing the leaves with the pestle using a circular motion until homogenized (about 1-3 minutes). Take the homogenate and filter through three layers of cheesecloth into a beaker pre-chilled on ice. The filtrate should be centrifuged at about 500 X g for 2–3 minutes to remove debris. The supernatant should be centrifuged at about 10,000 X g for 10 minutes to pellet the chloroplasts. Decant the supernatant and re-suspend the chloroplasts in a small volume of STK.

It is always crucial to assure that the centrifuge tubes are balanced. That is, the same weight in each of the two tubes that are placed in holders on opposite sides of the centrifuge rotor.

Below is a step-by-step flow chart for the isolation of chloroplasts from spinach:

Constructing a Flow Chart

While researchers rely upon written protocols similar to the one presented above, when working at the bench, they will usually prepare a flow chart similar to the one at right. You will find such flow charts helpful over the course of the term, especially as lab exercises become more complex.
  1. Prechill beakers, mortar and pestle, centrifuge tubes etc. on ice.
  2. Cut leaves into squares about 2 by 2 cm. Use scissors, and do it quickly.
  3. Place about 10 grams in the mortar.
  4. Add 50-100 ml of STK.
  5. Homogenize with a circular motion of the pestle.
  6. Filter homogenate through 3 layers of cheesecloth into a pre-chilled beaker.
    The following centrifugations should be performed at 4 degrees Celsius, and the final chloroplast suspension should be kept on ice.
  7. Centrifuge the filtrate at 500 X g for 2–3 minutes to remove plant debris.
  8. Centrifuge the supernatent at 10 000 X g for 10 minutes to pellet the chloroplasts.
  9. Decant the supernatent and resuspend the chloroplasts in a small volume of STK.
  10. View the chloroplasts under the microscope, using bright-field, phase constrast and fluorescence.

Chloroplast Preparation from Algae

When you harvest the algal cultures, the technique for isolating chloroplasts will vary depending on the type of algae. If you are provided with Chara australis, it is as simple as cutting one end of the internode cell (with small scissors) and squeezing the cell sap (with chloroplasts) onto a microscope slide.

Glass homogenizer.png
Eremosphaera viridis culture.png

If you are provided with Eremosphaera viridis, the large cells need to be broken apart. This is done with a glass mortar and pestle. The cells are decanted from the culture flask into a 50 ml disposable centrifuge tube, allowed to settle (this will take less than 5 minutes at "1 X g"). The supernatent (culture media) is carefully poured off (into the sink) and the cells re-suspended in STK. Then, they are poured into the glass homogenizer (Figure 1). The plunger is pushed to the bottom while twisting it. This is repeated 4 times, at which point the cells will have been disrupted, releasing the chloroplasts. The homogenate is filtered through multiple layers of cheesecloth, and the filtrate (chloroplast rich) is used as is for microscope visualization.


For all of the plant material, first observed the intact leaf or cell under the microscope, using bright-field, phase contrast, and fluorescence. Take careful note of the structure of the chloroplasts in situ. In the case of leaves, it is often useful to peel the epidermis off the leaf, and view the leaf with the peeled side up. On the microscope slide, samples of leaves can covered with pure water, and algal material can be covered with a drop of the culture medium. Chloroplasts should be covered with STK. Compare the intact leaves or cells with the isolated chloroplasts.

Under bright field illumination, you should be able to measure the size of chloroplasts in the various tissues (chloroplast will be identifiable by their ‘green’ color).

With phase contrast, you may be able to discern a phase halo around chloroplasts in situ. Compare this with the presence (or absence) of a strong halo around isolated chloroplasts. The phase halo is caused by the close apposition of two membranes in intact chloroplasts. Thylakoids will not have a strong halo since they have only one membrane. This is one way to assess the ‘intactness’ of the isolated chloroplasts. Fluorescence allows you to obtain direct evidence for the presence of chlorophyll, the only strongly fluorescent pigment in photosynthetic tissue.


How Fluorescence Microscopy Works

The technique used in microscopy is known as epi-fluorescence. Light of a wavelength suitable to excite a pigment (chlorophyll in our case) is transmitted through the objective with a dichroic filter --which reflects specific wavelengths, and allows others to pass through. The fluorescence light emitted from the pigment is focussed through the objective and selectively viewed with the emission filter (transmitting red light in the case of chlorophyll). Diagram re-drawn from Wikipedia: en.wikipedia.org/wiki/Fluorescence_microscope
Fluorescence Microscopy 01.png

With epi-fluorescence microscopy, you should be able to observe chlorophyll fluorescence in situ and in vitro. On the basis of your observations, compare the intactness of isolated chloroplasts with chloroplasts in situ.

For isolated chloroplasts, you simply take a drop of the suspension, place on a glass microscope slide and gently place a cover slip on top.

For in situ chloroplasts (in the leaf), the challenge is to slice through the leaf so that you will be able to see structures at the bottom and top of the leaf. One technique is to slice at a diagonal through the leaf. Another technique is to 'peel' the epidermis so that you can look directly at the internal architecture. Carefully place the leaf slices on a glass microscope slide, add a drop of water, and gently place a cover slip on top.



Hypotheses and Observations

The observations you make are qualitative in nature. For intact tissues and cells, how does the positioning and density of the chloroplasts relate to what you know about the spectral properties of intact leaves? Are chloroplasts dense enough? Would you predict 100% absorbance based upon their distribution in the leaf? For isolated chloroplasts, how do they compare with chloroplasts in situ? Would you predict that damage will affect the nature of the biochemical reactions the chloroplasts are capable of performing? For this question, take note that the light reactions are located on thylakoid membranes, carbon dioxide fixation and the Calvin cycle are located in the stroma of intact chloroplasts.

Personal tools
Namespaces
Variants
Actions
Navigation
Toolbox