The regulation of CF₁ is accomplished in two ways: 1) the proton gradient itself activates the enzyme; presumably, the proton "pressure" operating from the lumen of the thylakoid is sensed by the enzyme (probably by protonation of certain groups) (1); 2) light driven electron transport provides reducing equivalents that are used to reduce an intramolecular disulfide bridge (2) in CF₁, by way of a small protein, thioredoxin, which in turn is reduced by ferredoxin. Light thus provides the reducing equivalents and the proton "pressure" needed to activate CF₁. In darkness, the proton gradient disappears and CF₁ is oxidized to the disulfide state. These two effects interact so that reduced CF₁ is totally activated at a pH gradient of 2-3 units. It takes over 4 units to activate the oxidized form (3). If CF₁ can be reduced in the presence of a proton gradient, when there is no ADP or P_i to make ATP with, it can act as an ATPase, hydrolyzing available ATP. This hydrolysis, however, is accompanied by proton pumping to the interior of the thylakoid, so illuminated thylakoids should be much less effective at ATP hydrolysis (because light is already establishing a proton gradient).

If Mitchell is correct, that a proton gradient is the high energy "intermediate" driving ATP synthesis, then we should be able to bypass light absorption and electron transport completely and make ATP by artificially producing an environment in which the interior of the thylakoid is buffered at a low pH and the exterior is buffered at a high pH. It would be interesting to test a range of pH gradients (with or without DTT - remember activation) to measure their effectiveness in ATP synthesis. Is there some threshhold DpH below which no ATP can be made?

One of the requirements in scientific work is reproducibility. When you design your experiments, keep this goal in mind. Replicate points, especially references. Think about the order of addition of substances. Think about possible controls. How will you measure the amount of ATP produced? How will your controls verify that this method is working? For example, it might be a good idea to add a known amount of ATP to a "dummy" tube to validate your ATP measurement. Any experiment should be done at least twice to verify that the results are "real". If an experiment does not produce the result you expect, check through your protocol to see if you have left something out or if there is a flaw in your design. Don't repeat a failed experiment blindly. Have some reason for doing so, some change in the protocol or in your skill level that makes success more likely.

When presenting your results, do so in a way that makes them comparable with others. Generally, you do not present raw data. Rates are normalized to some measure of the amount of "stuff" catalyzing the reaction - mg of protein or chlorophyll, for instance. Time courses should generally be plotted. Standard curves do not need to be presented in a report, unless there is something unusual about them. Replicate measurements should not be averaged, unless there are several that are in good agreement; if an average is given, a standard error or deviation must be also. Be honest about discarded data; give a reason for discarding it.

There are several questions you can pursue in this lab. Can we measure photophosphorylation (ATP synthesis) this way? What compounds can serve as electron acceptors? Is there a difference between electron acceptors in terms of the efficiency of phosphorylation? What sorts of compounds inhibit photophosphorylation (other electron acceptors? uncouplers? etc.)? How does photophosphorylation depend on the wavelength of the light? Will an artificial proton gradient drive ATP synthesis in the absence of light? To what extent? How large a gradient is needed? How long does it take for a buffer to permeate to the inside of the thylakoid? Will the same inhibitors work in ATP hydrolysis as worked in photophosphorylation? Does the presence of DTT make a difference? Under what conditions will thylakoid-bound CF₁ hydrolyze ATP? Might there be other ways of activating the CF₁? Will the same inhibitors work here? How does CF₁ behave when not bound to the thylakoid? Can it make ATP? Does it need to be activated? What will activate it when off the membrane? Will the same inhibitors work? These are some of the questions you might wish to pursue.

These questions require some kind of baseline measurements for comparison purposes. In order for these comparisons to be meaningful, you have to be confident of your baselines. For instance, if you are measuring the rate of photophosphorylation under different conditions, it is important to have some standard conditions for which you know the rate quite accurately. This means you will need at least two good measurements of this rate. The same thing goes for ATP hydrolysis rates. It is best to check the photophosphorylation of your thylakoids every day, since individual preparations may vary.

Stability of reagents. Buffers should be kept in the cold. Enzyme solutions are generally stable at 4° for a day. If dissolved in a glycerol containing buffer, or stored as a precipitate, they will generally be stable for weeks. ATP and ADP are stable in cold or frozen solution for a long time if the pH is near 7. Electron acceptors are generally light sensitive. NAD⁺ solutions are stable frozen. Chloroplasts need to be made fresh each day they are used. They will be stable during the day if kept on ice.

Chloroplast preparation. Add 200 mL grinding buffer (0.33 M sorbitol, 40 mM bicine, 5 mM MgCl₂, 0.5 mg/mL BSA, 2 mM ascorbic acid, pH 8.0) to 40-70 g deveined, washed dark green leaves. Blend at moderate speed for the minimum time required to reduce the leaves to fragments of approximately 5 mm (5-10 s). Pour the mixture through Miracloth and sediment the chloroplasts at 3000 g for 1 min. Discard the supernatant and gently dislodge the pellet in 3 mL lysis medium (20 mM KCl, 20 mM NaCl, 5 mM MgCl₂, 25 mM bicine, 1 mg/mL BSA, pH 8.0). Using a Teflon pestle, homogenize the pellet in a cool 50 mL round bottom centrifuge tube. Bring the volume to approximately 45 mL by addition of lysis medium, and resediment at 7000 g for 5 min. Discard the supernatant and resuspend the pellet as before in approximately 5 mL resuspension buffer (0.33 M sorbitol, 5 mM MgCl₂, 10 mM NaCl, 25 mM bicine, 3 mg/mL BSA, pH 8.0). Store on ice and in the dark. There should be approximately 10 mg of chlorophyll in this sample.

Chlorophyll determination. Add 4 mL 80% acetone to four glass tubes. To two, add 20 mL of your chloroplast suspension; to the other two, add 40 m L. Mix vigorously. Sediment in a benchtop centrifuge (5 min). Read the absorbance at 652 nm in a glass cuvette. Use 80% acetone as the blank or reference. The chlorophyll concentration in mg/mL is given by multiplying the A₆₅₂ by 2.8 for the 40 mL samples, or by 5.6 for the 20 mL samples.

Photophosphorylation. Isolated thylakoids have lost ferredoxin, and so have no place to dispose of the electrons generated by photosystem I. It is necessary to add an electron acceptor to take ferredoxin's place. Two that have been used frequently are PMS (N-methylphenizinium methosulfate) and pyocyanine. PMS will cause problems later on in the determination of ATP amounts. Having called all over the country, I am pretty sure that pyocyanine is no longer commercially available. It will be interesting to try other possible acceptors (chlorpromazine, ferricyanide, methyl viologen, anthraquinone 2-sulfate, methylene blue are some), to see if they will sustain efficient photophosphorylation. PMS destroys NADH produced from glucose 6-phosphate in the reactions done after photophosphorylation (see below). We can protect the NADH briefly with a high concentration of DTT (a reductant). Chlorpromazine and methyl viologen are harmless to NADH, at least in the short run. Other potential electron acceptors will have to be tested. DCMU is known to interfere with chloroplast electron transport. You might want to study its effects.

In order to avoid problems with ATPases that may be present in the thylakoids (including CF₁), we will use a "hexokinase trap" to maintain [ATP] at very low levels. As the ATP is made, hexokinase will transfer the phosphate to glucose to make glucose 6-phosphate. Thus, the ATP is "trapped" in the glucose 6-phosphate. We can then measure the glucose 6-phosphate at a later time.

To measure the time course of ATP production, we need to be able to stop the reaction quickly. Since we have to do a further enzyme catalyzed reaction to measure the glucose 6-phosphate, we want to stop it in a way that will be harmless to this later reaction. NH₄⁺ dissipates the proton gradient quickly, and so should stop the reaction. We will use NH₄Cl as a source of ammonium ion. (Could this be used as an inhibitor?)

To measure photophosphorylation, add chloroplasts (final concentration 25-50 mg chlorophyll/mL) to a solution of 25 mM bicine, 50 mM KCl, 5 mM MgCl₂, 1 mM ADP, 3 mM NaP_i, 10 mM glucose, pH 8.0, 10 units/mL hexokinase and 25 to 50 mM electron acceptor (I'm not sure about ferricyanide). Hexokinase and electron acceptor should be made up as separate solutions and added immediately prior to the experiment. The last addition should be the chloroplasts. Illuminate immediately with a strong light source (like a slide projector), mixing constantly. Use a clear vial and stir with a magnetic flea bar. For each time point, withdraw 0.1 mL and mix immediately with 1.1 mL 25 mM bicine, 50 mM KCl, 5 mM MgCl₂, 10 mM NH₄Cl, pH 8.0, containing 0.5 units/mL glucose 6-phosphate dehydrogenase and 1 mM NAD⁺ added just before you begin the experiment. Measure A₃₄₀ to determine [NADH] (the molar extinction coefficient (absorbance of a 1 M solution in a 1 cm cuvette) for NADH at 340 nm is 6220 M^-1). If you are using PMS (or something else that destroys NADH) as an electron acceptor, you must add 5-25 mM DTT to the stop solution (the one with the NH₄Cl) and leave out the NAD⁺, to avoid having the NADH present for an extended time with the PMS. In this case, put your 1.2 mL in a cuvette, add NAD⁺ to 1 mM, mix rapidly and follow the time course of A₃₄₀. It should rise and fall over a few minutes. Take the maximum as a measure of NADH production.

For your zero time point, either remove the 0.1 mL immediately, or you can remove the 0.1 mL before adding the chloroplasts, and add 2.5-5.0 mg (1/10 of what you would add to 1 mL reaction mix) chlorophyll to that; then follow the regular workup. Notice that you need one mL of reaction volume for about 10 aliquots (including references) of this reaction mixture. You should also include a reference with a known [ATP] as an internal standard. This means you need to add a known amount of ATP to your photophosphorylation mix under conditions where the thylakoids cannot make ATP, but the hexokinase trap can still work. You want to be certain that your ATP detection method is working.

ATP synthesis without light. In this case, we wish to establish the DpH without benefit of light, so these reactions are carried out in the dark (non-illuminated). This raises the problem of activation, but since we will establish the gradient, and we can add DTT, CF₁ should be activated. To do this measurement, pellet a known quantity of chlorophyll in a microcentrifuge tube. Resuspend the chloroplast pellet in 10 mM NaCl to a concentration of 0.5 to 1 mg chlorophyll/mL. Mix this suspension with an equal volume of 10 mM succinate, 10 mM MgCl₂, pH 3.8. After five minutes on ice (you can try varying this time), add a volume of 0.1 M bicine, 5 mM MgCl₂, 1 mM ADP, 10 mM NaP_i, 20 mM glucose, pH 8.2 equal to the combined volume of resuspended chloroplasts and succinate buffer (your final reaction mix thus contains 1 part chloroplasts in 10 mM NaCl, 1 part succinate buffer, and 2 parts bicine buffer). Remove aliquots into microcentrifuge tubes and spin down quickly to remove thylakoids. You might want to try quenching in NH₄Cl as above. You will want to sample this reaction at earlier times (as early as 10 s) because there is nothing replenishing the proton gradient, so the rate of synthesis should drop fairly quickly. You can try adding DTT to any or all of the three solutions you use. It should be in the range of 5-10 mM.

Measurement of ATP:

Because of the small amount of ATP made, you cannot use the method described above. There are two other methods for measuring ATP that are more sensitive. One is to follow the protocol above and then use NADH fluorescence, rather than absorbance. This technique is roughly an order of magnitude more sensitive, with detection down to about 1 nmol.

Another alternative is to use the luciferin-luciferase method of measurement. This technique is capable of measuring quantities of ATP in the range of 1 to 20 pmoles. Firefly lantern extract (FLE) is dissolved in 25 mM bicine, 10 mM MgSO₄ at 10 mg/mL. Each assay requires 0.1 mL of FLE, so make up just enough. If kept cold (not frozen!) the solution can be used the next day, but the technique will be slightly less sensitive. 0.1 mL FLE solution is added to a 1 dram (or smaller) vial (these vials must be clean and kept in the dark for at least 24 hours). The vial is placed in a scintillation counter in photon counting mode to record the background light emission. When this level is established (2-10 min), the vial is removed (the room must be kept dark) and the solution to be assayed is added (1-20 mL). Mix for 10 sec by gentle shaking. Place vial in counter. Begin counting at an exact and reproducible interval after the addition of the ATP sample (15 or 20 sec works well). Count light emissions for some fixed interval; 2 or 3 minutes works well. Keep all solutions cold during this assay. You must produce a standard curve each time you do this assay. A 1 mM solution of ATP works well for this. Do at least three different amounts of ATP, in duplicate, for the standard curve.

This is technically the most difficult procedure to do successfully. For that reason, please do not select this as your principal or only project.

ATP hydrolysis. Since we will be measuring the reaction in the opposite direction, we need some other solutions:

Buffer A: 25 mM bicine, 50 mM KCl, 5 mM MgCl₂, set to pH 8.0 with KOH.

Buffer D: 50 mM bicine, 50 mM MgCl₂, 50 mM ATP, pH 8.0.

Phosphate stain: To 0.5 M H₂SO₄ add FeSO₄ to 4%. After it dissolves, add ammonium molybdate to 1%.

Because this is not an energy requiring reaction, it is of less interest to determine its time course (although you may want to). Our main interest here is to try different methods of activation (and combinations of methods) and determine their effects on ATP hydrolysis. The two obvious ones are illumination and DTT treatment. Illumination should be for a period of about 5 min (you can certainly vary this). DTT should be in the range of 5-10 mM (although you could also vary this). You could certainly try a pH gradient as well. In any case, you will need a control (with no activation). You must not add the ATP until the activation is complete. Because the reaction requires no energy, we cannot use NH₄Cl to stop it. We will use 10% perchloric acid (PCA) instead. Illumination will probably require an electron acceptor to work (you might test this, also) since its function is to establish an acidic interior. Since we are not using NADH here, you can use PMS without worry. Put 0.9 mL buffer A in a glass tube and add electron acceptor. If you are using a chemical activator, add that now as well. Then add 25 mg chlorophyll and incubate (remember, for about 5 min) with the activator. Then add 0.1 mL buffer D (do NOT make more of buffer D than you need - it takes a lot of ATP) and mix well. Don't forget to prepare a reference tube (not just a control with no activation, but a reference in which the reaction is stopped before ATP is added). After a defined time interval (usually 2-4 min, use the same for each sample), add 0.25 mL 10% PCA to stop the reaction. Mix well and keep on ice. For illuminated samples, add buffer D just before switching off the lamp. DO NOT illuminate during the ATP hydrolysis. Centrifuge your samples to sediment the denatured chloroplasts. Withdraw 1 mL of the supernatant, and add to 1 mL of the phosphate stain. Mix well and incubate for 5 min. Then add 0.2 mL 1 M citric acid to stop color development and measure the absorbance at 700 nm. At the same time, you need to prepare a set of P_i standards in the range of 0.1-1.0 mM. These need to be treated in essentially the same way as your experimental tubes. (1 mM P_i should give an absorbance of about 1.4).

CF₁ activity away from the membrane. Of course, the first order of business is to separate the CF₁ from the membrane. There are a number of ways to do this. We will use chloroform to effect the separation.

Buffer F: 0.33 M sorbitol, 20 mM Tris, 1 mM EDTA, pH to 7.6 with sulfuric acid.

Buffer G: 25 mM bicine, 5 mM CaCl₂, 5 mM ATP, pH 8.0.

CF₁ is cold-labile, so these steps must take place at room temperature. Add 1-2 mL chloroplasts to 40 mL buffer F (these chloroplasts may be up to a day old - we don't care about coupling here). Mix and centrifuge at 7500 g for 5 min. Resuspend the pellet in 0.5-1.0 mL buffer F and keep in a microcentrifuge tube. Adjust total volume to about 1.2 mL. Determine the chlorophyll concentration (as above). Add 30 mL chloroform per mg chlorophyll and vortex for 25 s. Sediment the resulting debris in a microcentrifuge for 15 min. Withdraw and keep as much of the clear supernatant as you can without disturbing the pellet (should be at least 1 mL). Add sufficient ATP (from a 0.2 M solution) to bring to 2 mM. Store at room temperature.

Use a standard method to determine the protein concentration. It is likely to be 0.2-1 mg/mL.

The rest of this experiment is similar to the ATP hydrolysis experiment above. CF₁ will not synthesize ATP away from the membrane (although you could verify this, if you wish). It is more convenient to activate in a smaller volume, since the CF₁ solution already contains ATP. Activate 0.2 mL CF₁ in a test tube by adding DTT, or heating to 60^o (or some other possible activation scheme), for 5 min. If heating, place on ice immediately for 60 s. and then keep at room temperature (capped). Remember to try combination(s) of activators as above. Also, remember to include controls and references. To measure ATP hydrolysis, add 10 mg CF₁ to 1 mL buffer G (you might want to try other divalent cations besides Ca⁺²) and incubate at room temperature for 10 min. Stop by adding 0.25 mL 10% PCA. After mixing, add 1.25 mL phosphate stain and mix. After 5 min, add 0.2 mL 1 M citric acid to stop color development and measure the absorbance at 700 nm. You may have to adjust CF₁ amounts or incubation times to get readable samples. Remember to do standards, as above.

You might want to examine the effects of any inhibitors on this reaction.

Calculating results.

I will give you one example. Let us say that I have done the photophosphorylation experiment, and at 5 min, my sample shows an A₃₄₀ of 0.12. This corresponds to [NADH] @ 0.02 mM. Since I measured the NADH in 1.2 mL, I have 1.2 x 0.02 = 0.024 mmol NADH, indicating that 0.024 mmol ATP was produced, for an apparent rate of 4.8 nmol/min. If I added 40 mg chlorophyll to my initial reaction mixture, my apparent rate of synthesis, normalized for chlorophyll content, is 4.8 nmol/min/4 mg chlorophyll = 1.2 nmol/min-mg chlorophyll (my aliquot had 1/10 of the total chlorophyll).

1. Mills, J.D. and Mitchell, P. (1984) Biochim. Biophys. Acta 764, 93-104.

2. Ketcham, S.R., Davenport, J.W., Warnke, K. and McCarty, R.E. (1984) J. Biol. Chem. 259, 7286-7293.

3. Junesch, U. and Gräber, P. (1987) Biochim. Biophys. Acta 893, 275-288.