Rubisco may already be familiar to you from Cell & Molecular Biology. Now you will have a chance to investigate the enzyme itself a little more thoroughly. It is important as you begin that you have some questions in mind that you would like to answer. For instance, you might want to determine Rubisco's relative activity with CO₂ vs. O₂. Rubisco as isolated usually is not very active, but requires a covalent modification to become active. You might want to examine the conditions for activation and/or inactivation. You have been told that the large subunits are the catalytic subunits. Can this be verified? Can you get some idea of the role of the small subunits? Can you even separate the subunits, without destroying the protein's 3° structure? Can you use a gel to accurately determine the MW? Are Rubiscos from different sources similar in MW? And so on...

There are two approaches to the initial isolation of Rubisco. In one, chloroplasts are first isolated and then lysed by hypotonic treatment. This is the procedure used in Cell and Molecular Biology. In the other, the plant cells are thoroughly macerated and the cytoplasm/stroma mix is treated to isolate Rubisco. You will use the second method.

Initial Rubisco isolation

Wash and devein 50 g of leaves. Add 125 mL cold bicine buffer (25 mM bicine, 1 mM EDTA, pH 7.8), containing 2% PVP (polyvinylpyrrolidone). In a blender, blend in short bursts until all the leaves are fragmented. Then blend 5-8 s on high. Filter through 8 layers muslin, followed by filtration through 2 layers miracloth (or muslin).

Spin at 10-11,000 g for 30-40 min. Carefully remove the supernatant into a graduated cylinder and note the volume. Transfer to a beaker and, while stirring, add 3/7 volume 60% PEG (MW 3400) to bring to 18% PEG. Stir in the cold for 20 min. Centrifuge at 24,000 g for 20 min. Pour the supernatant into a graduated cylinder. Note the volume. Transfer to a beaker and, while stirring, add 0.01 volume 2 M MgCl₂ to bring the final concentration of Mg⁺² to 20 mM. Stir 5 min in the cold. Centrifuge as before. Resuspend the pellet in no more than 1 mL bicine buffer (w/o PVP). Clarify by spinning at 14000 rpm for 15 min. This supernatant contains the isolated Rubisco and can be used for assays, gels, etc., or further purified by column chromatography (below).

Assay for Rubisco activity

An assay is a means of measuring the amount of catalyst (active enzyme) present. Conditions are chosen such that the catalyst is operating at V_max, or well over the K_m. At this concentration of substrate(s), the velocity is just k_cat[Enzyme]_total. Since k_cat is the same for all rubisco molecules, the velocity is a measure of the enzyme concentration. Thus, even in a mixture, we can measure the amount of a particular enzyme that is present if we can develop reaction conditions that are specific to it.

This is a coupled or linked assay, in which the immediate product of Rubisco, 3-phosphoglycerate, is converted first to 1,3- bisphosphoglycerate, and then to glyceraldehyde 3-phosphate using ATP and NADH, and the enzymes, phosphoglycerokinase and glyceraldehyde 3-phosphate dehydrogenase (glycolytic enzymes). We follow the disappearance of NADH by monitoring the A₃₄₀. The extinction coefficient at this wavelength is 6.22 mM^-1 cm^-1.

There are a number of cautions that need to be considered in this assay. First, keep everything cold until the assay. This is especially true for enzyme solutions and for solutions containing labile compounds, such as ATP, NADH and ribulose bisphosphate. Second, Rubisco usually requires "activation" in bicarbonate. This means that you must add your sample to the reaction buffer and let it sit several minutes before adding substrate (RBP). Third, RBP is only available in limited quantities - do not make more than you need. Finally, in all spectrophotometric assays, adding and mixing are critical. Remember that you want to measure initial velocities. Once the reaction begins you need to begin measurement as soon as possible. On the other hand, your data will not be useful if the solution is not well mixed. You must mix quickly and effectively. This usually takes some practice. Be prepared to "waste" some assays.

Procedure

There are several components to the reaction mixture. These are:

sample: your protein sample

water: to make up the total volume.

 
Ion exchange chromatography

Important: if you are not going to do the column chromatography immediately following the initial isolation, it is best to keep the rubisco in its precipitated form (after MgCl₂ addition) in the cold. Immediately prior to running the column, collect the precipitate by centrifugation and resuspend and clarify as described above. Save a small volume of the resuspended pellet for assay along with column fractions.

Final protein purification frequently involves ion exchange chromatography. The most common resin used for proteins is DEAE- cellulose. This is just cellulose derivatized with an ethylamino group, that has two more ethyl groups attached to the nitrogen. Like most amines, this group has a pK_a of about 9, so at neutral pH, it is positively charged. The average charge on the group can be manipulated by changing the pH. Most proteins have a negative charge at neutral pH, so they bind to the DEAE group.

Most ion exchange chromatography requires washing the column with a gradient of salt. That is, you begin washing with a low ionic strength buffer, and gradually increase the ionic strength by the addition of salt. Different proteins bind to the DEAE group more or less tightly, and so are removed from the column at different ionic strengths. "Ion exchange" refers to the process of exchanging the protein for some other ion binding to the DEAE.

In practice, this gradient is achieved by having two beakers or flasks, one containing the low ionic strength buffer and one containing the buffer with as much salt added as you wish to have at maximum ionic strength (usually 0.2 - 0.5 M). The column is fed from the first beaker. The two beakers are connected by gravity feed. As the level in the first beaker drops, solution from the second feeds into it, thus raising its ionic strength. The effect is to generate a washing solution of slowly increasing ionic strength.

In this case, we will use 50 mM bicine, 0.1 mM EDTA, 5 mM DTT, pH 8.0 as the column buffer. Be sure to add the DTT after setting the pH. The high ionic strength solution will contain column buffer, plus 0.5 M NaHCO₃, at the same pH. Before you run the column, it must be equilibrated with the column buffer, which means you need to run 200 mL of buffer through the column. After you run the column, you need to clean it off with 200 mL of buffer containing 0.6 M salt (NaCl or KCl will do).

Let the buffer on top of the column run through until none is left. Add your protein sample and let it run into the column. Then wash the column with 100-200 mL column buffer (without bicarbonate). Now you are ready to run the gradient and collect the eluant.

Material leaving the column is collected in test tubes by a fraction collector (usually inanimate). You will want to collect about 500 mL (the gradient) so set your fraction volume accordingly. The absorbance of these fractions at 280 nm is determined, as an indication of protein content. Then, a search is made through the tubes for enzyme activity. Usually this is confined to a few fractions (<10). These fractions are pooled and precipitated by the addition of (NH₄)₂SO₄ to a final concentration of 50% saturation (add 0.3 g/mL of solution). The protein is stored in this form. It is best to do the activity assays the same day as the column.

Lowry assay and [protein] determination

Before running SDS PAGE, you should know how much protein there is in each sample. It is important to both load enough protein on the gel to be visible, and to load the lanes equally.

You will determine protein concentration by using the Lowry procedure. You will prepare two tubes for each sample, as well as two tubes for each concentration used on the standard curve. Each tube will contain a protein sample plus enough water to bring the final volume to 0.5 mL. You must do a standard curve each time you do a Lowry assay. This curve is generated using a BSA (bovine serum albumin) solution of known concentration (0.2 mg/mL). You will have six points on the standard curve.

water	BSA
0.5 mL	0
0.475	25 mL
0.45	50
0.4	0.1 mL
0.25	0.25
0	0.5

In general, add the water first, and then the BSA. This will give you 10 tubes containing BSA (5, 10, 20, 50 and 100 m g, respectively). The rest of your tubes will be your samples. Some samples will be fairly concentrated, so you should do a dilution for the protein assay. Some samples may be very dilute, so you will need to add more for them. Once you have decided what volume of sample you will be using, add enough water so that the total volume will be 0.5 mL. Then add the sample (you should use volumes between 10 and 500 m L). After you have prepared all the standards and samples, add 1.5 mL Lowry D solution to each and vortex. Let stand for 10 min. Add 0.15 mL E to each and vortex immediately. Let stand in the dark for 30 min. and read the absorbance at 750 nm. Samples that have a lot of chlorophyll in them may give abnormally high absorbances. To control for this, put the same volume of sample used in the Lowry, above, into a test tube. Add enough water to bring the total volume to 2.15 mL. Read the absorbance at 750 nm. Subtract this absorbance from the absorbance of your Lowry sample.

Notes on Lowry procedure: The Lowry is what is known as a colorimetric assay. That is, a color is developed whose intensity is proportional to the concentration of the substance of interest, in this case, protein. The more protein, the bluer the test tube should be after the half hour incubation in the dark. When you have the absorbances of blank, standards and samples, you should plot the absorbance vs. the amount of protein in mg for the standards. Then draw a line through the points. From this graph you will be able to determine your protein concentrations in each sample.

Lowry solutions

A 2% Na₂CO₃, 1% SDS, 0.16% K-Na tartarate in 0.1 M NaOH

B 4% CuSO₄^.5H₂O

D 100 mL A, 1 mL B

E 1 M Folin Phenol Reagent (1:1 dilution with H₂O)

Once you have the concentrations of your protein samples, you are ready to run the SDS PAGE to determine the relative amounts of Rubisco in each sample.

Gel electrophoresis

Preparation of samples for SDS PAGE:

Once the gel is poured, the gel sandwich is placed in the apparatus and held in place with the binder clips. The top junction between the glass plate and the apparatus must be sealed with melted agar. Add electrode buffer to the reservoirs on the top and on the bottom. Remove air bubbles from the bottom of the gel by tilting it back and forth, before sealing. The gel should be run in the cold room.

Samples were already prepared by boiling with sample buffer. Immediately before adding them to the gel, boil them again for 2- 3 min. You will want to add 30-50 m g of protein to each well. Try to apply the same amount of protein from each sample. If unsure of the [protein] add the sample to two lanes in different amounts. Apply your samples, being careful to note which sample is placed in which lane. Attach the cables to the apparatus, with the red cable being attached to the bottom electrode (this is the anode). Turn on the power supply. The samples should run through the stacking gel at about 80 volts. Once into the running gel, increase the voltage to 160 volts to complete the run. It should take about 2-3 hr to run the gel at these settings. Turn off the power supply and turn the controls to zero when the tracking dye is with 1 cm of the bottom of the gel.

Staining the gel

This is a very easy procedure. The hard part is getting the gel out of the glass plates.

Using a plastic spacer, gently pry the top plate off of the gel. Carefully lift the gel off the bottom plate. A little water may help, here. Place the gel in a glass or plastic dish whose bottom is not much larger than the gel. Stain the gel with Coomassie blue. Cover with stain for 1-2 hours, with shaking. Then pour off the stain, cover with the destain solution for another 2-4 hours (more is all right). The destain solution must be changed periodically, probably 3 or 4 times. Continue to destain until the background is nearly clear and blue bands from the proteins are clearly visible.

Running gel

For 8 mL:
 

H2O	2.82 mL
1.5 M Tris pH 8.8	2
40% acrylamide	2
2% bis-acrylamide	1.1
TEMED	4 mL
10% SDS	80 mL
(or 8 mg solid SDS)

Mix well. While mixing, dissolve 50 mg ammonium persulfate in 0.5 mL water. Add 18 m L APS to the gel mixture, stir for about 1 min. Make sure it is well mixed, but do not create a lot of bubbling. Quickly pour the gel into the sandwich to fill as described. Carefully add a small amount of butanol to the top of the gel. You should see a straight line marking the interface between the gel and the butanol. When the gel has polymerized, pour off the butanol.

Stacking gel

For 4 mL:
 

H2O	1.91 mL
0.5 M Tris pH 6.8	1
40% acrylamide	0.45
2% bis-acrylamide	0.6
TEMED	4 mL
10% SDS	40 mL
(or 4 mg solid SDS)

Mix as before. Add 8 mL APS, mix again, and pour enough to fill about ¾ of the remaining volume in the sandwich. Add the comb, and finish filling.

Electrode buffer

3.03 g Tris base, 14.4 g glycine, 1 g SDS, in 1 L H₂O.