Pyrene fluorescence assay of actin assembly II

Dyche Mullins, 9/13/99

Stock solutions:

Buffer A (a.k.a. G buffer):
2 mM Tris HCl, pH 8.0
0.2 mM ATP
0.1 mM CaCl2
0.5 mM DTT

KMEI (Polymerization buffer):
50 mM KCl
1 mM MgCl2
1 mM EGTA
10 mM Imidazole HCl, pH 7.0

ME (Mg-exchange buffer):
50 µM MgCl2
0.2 mM EGTA

Useful facts:

Extinction coeff of actin at 290 nm: 38.5µM/O.D.
Correction for pyrene actin - subtract 0.127´A344 from A290
Extinction coeff of pyrene at 344 nm: 45.0µM/O.D.

PREFERRED WAVELENGTHS:
Excitation: 365 nm
Emission: 407 nm

ACCEPTABLE WAVELENGTHS:
Excitation: 365 nm
Emission: 385 nm

Before starting:

1. Be sure you have good reagents.
a. Actin must be gel-filtered and you should only use fractions from the back side of the gel filtration peak.
b. You can use 100% pyrene labeled actin but be careful. Not all actin-binding proteins bind as tightly to pyrene actin (e.g. profilin!). The perferred range of pyrene doping is 1-10%.
c. Don't use frozen actin for nucleation assays, it assembles with measurably slower kinetics.
d. Don't use old actin. Monomeric actin doesn't store very well. If you're particularly conservative don't use actin more than two weeks old. If you're not so conservative you can use it for 4-6 weeks but be aware that you are pushing your luck. Pyrene actin keeps better than dark actin so you can use quite old (several months old) pyrene actin with fresh dark actin.
e. Check your polymerization curves versus a standard curve. If you have an untried batch of actin, do a spontaneous polymerization assay at a known concentration and compare it to the curves attached to this sheet. If there is a big difference then either i) your concentration is off or ii) your actin is bad. Do not proceed until you have figured this out. Wasting time is bad!
2. Turn on the fluorimeter and fire up the computer. Make sure everything is turned off at the beginning. The correct sequence of events for the ISS K2 is:
a. Turn on the arc lamp - This is the important part. The lamp is a 300 W xenon arc lamp and it can kill nearby electronic equipment and unsupervised children. Turn the current control knob all the way to the left (minimum current) and flip the switch. Wait until you hear the audible pop of the arc starting and/or until you see the panel meter read 10 Amps.
b. Turn on the fluorimeter. The switch is in the right rear of the instrument. Make sure the PMT you want is selected and set to the correct voltage. For most applications, use the PMT on the right hand side. Set the toggle on the PMT to "manual" and dial up a voltage between 900-970V.
c. Turn on the computer. Make sure to switch on both the monitor and CPU. Wait for the computer to boot up and then, at the prompt, type "iss". For polymerization assays use the "Take a Spectrum" option (#2). Then choose "Intensity" (option #2 again) and "Slow Kinetics" (also option #2).
d. Finally bring the lamp up to operating voltage. Slowly turn the current knob until the panel meter reads 15 Amps.
e. The shut-down procedure is the exact reverse of the above sequence.
3. Optimize the fluorimeter. You want to maximize signal and minimize light scattering and photobleaching.
a. Photobleaching. To minimize photobleaching, you must minimize the excitation light. Settings that have worked for me are: 0.5 mm excitation slits, excitation polarizer in line (this also seems to help with light scattering) and excitation iris closed 80-90%. Remember, you cannot simply correct for photobleaching mathematically. There are free radicals involved. Very dangerous.
b. Light scattering. Large structures (aggregates, vesicles etc.) scatter incident light. This causes a problem because the monochrometers on the spec aren't perfect and will pass a small amount of light at the excitation frequency. You can cut down on light scattering by spinning and degassing your sample before use, by putting the excitation polarizer in line (the monochrometers are more selective when dealing with vertically polarized light), by using the smallest monochrometer slits possible, and by using a low-pass filter on the emission channel (the 399 nm cutoff works well for pyrene fluorescence).
c. Signal. If you don't have enough signal, or if your signal to noise ratio (SNR) is crappy, you can do several things: i) increase the ratio of pyrene:dark actin, ii) increase the emission slit width, iii) increase the PMT voltage (don't go up to 1000 volts), iv) increase the excitation light by opening the iris or increasing the slit widths or v) as a desperate measure, you can use the left emission channel (this channel has no monochrometer so the light throughput is much higher. You just need a good low pass cutoff filter).
d. Check your settings! Polymerize 100 uL of your stock solution of pyrene actin then dilute it down to working concentration and load it into the fluorimeter. Collect data for several hundred seconds. If the signal is trending downward, you are photobleaching and you should reduce your excitation light. If your signal is too strong you should also decrease the size of the excitation and/or emission slits. If the signal is too weak, see step (c) above.

Tips for collecting good data:

1. Polymerize by adding 0.1 volumes of 10X KMEI. This buffer gives the optimum polymerization rate (See Drenkenham and Pollard, 1984 J. Biol. Chem. for details) and also approximates the intracellular ionic composition. Bring all of your stock solutions together and triturate several times for good mixing. Schlieren lines are your enemy!
2. Pre-exchange the calcium for magnesium. Monomeric actin is stored in calcium and if you polymerize it by simply adding 10X KMEI, the Ca-Mg exchange shows up in the kinetics. To pre-exchange the Mg, incubate the pyrene-doped actin stock solution with 0.1 vol. of 10X ME (see above) for about 2 minutes before adding the KMEI. Do not pre-exchange all of your actin at once. It will polymerize before you can do the assays!
3. Keep your buffers at room temperature. If your sample starts out cold at time zero, and warms to room temperature during data collection, the kinetics will be way off. If you are adding a large volume of buffer to small volumes of protein (say 10:1), you can keep the protein on ice and the buffer at R.T.
4. Keep track of the dead time. The ISS does an admirable but lamentably slow dark count correction for each data set. To collect analyzable data use a stop watch. Start it as soon as you begin the polymerization reaction and stop it as soon as the first data point shows up on the screen. Record the dead time for each data set and add it back to the X-axis data before analysis. In this manner you may more quickly approach the way of the kinetic warrior.
5. Watch for air bubbles. When loading, tilt the 60 µL quartz cuvette to one side and put your pipette tip in one corner. Fill slowly and watch for bubbles. If you get a bubble, withdraw the liquid and try again. As long as you're measuring dead time with a stopwatch it's okay. Take a deep breath. Try again. Bubbles kill data dead!
6. Use a fresh box of pipette tips and keep the lid closed when not in use. Dust is bad.
7. Wash the cuvette thoroughly between each run. Little bits of leftover actin will nucleate filament formation in your next run. Use the cuvette washer and don't spare the water. Rinse water, ethanol, water - both inside and out. Wipe down the outside to remove fingerprints. Dry the cuvette on the washer before reuse.
8. Collect complete data sets from time zero until the pyrene fluorescence plateaus. You cannot compare or analyze partial data sets.
9. If you have determined that the conditions you are using don't alter the critical concentration, you can normalize your data. First, subtract the baseline values (fluorescence at time zero) from each data set. Then divide all the data in each dataset by the fluorescence at the plateau. This will correct for small pipetting errors and variations in the cuvette position.
10. Make your mind right. Remember what Henry V says: "All else be so, if our minds be right."