Pyrene Fluorescence Assay

Pyrene Fluorescence Assay of Actin Assembly

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
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 (low light scattering):
Excitation: 365 nm
Emission: 407 nm

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 previously 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 diluted into 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 below. If there is a big difference then either i) the actin concentration is wrong or ii) the actin is bad. Do not proceed until you have figured this out.

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 most fluorimeters is:
a. Turn on the arc lamp – This is the important part. The lamps on most fluorimeters are 200-400W xenon arc lamps and they can put a spike on the power line that will damage sensitive electronic equipment. Set the current at its minimum value.

b. Turn on the fluorimeter. This includes photomultiplier(PMT) power supply and monochrometer motors. Select an appropriate PMT voltage.

c. Turn on the computer.

d. Finally bring the lamp up to its appropriate operating current. The optimal value varies from manufacturer to manufacturer. Low lamp currents can cause unstable arcs and unacceptably noisy data so it is usually best to find the optimal lamp current and then vary the excitation intensity with slits and filters (we had a significant noise problem with our ISS fluorimeter before we figured this out).

e. The shut-down procedure is the exact reverse of the above sequence.

3. Optimize the fluorimeter settings. You want to maximize signal and minimize both light scattering and photobleaching. Start by polymerizing pyrene actin at the concentration you will be using in your experiments. Place this solution in the fluorimeter and use its fluorescence to help in adjusting the settings. For best results, proceed in the following order:
a. Light scattering. Large structures (protein aggregates, lipid vesicles etc.) scatter incident light. This causes a problem because the monochrometers 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 the sample before use, by putting a polarizing filter in front of the excitation light (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 (a KV399 Schott glass filter works well for pyrene fluorescence).

b. Photobleaching. If the fluorescence signal at steady state is a flat line, you’re okay. If the fluorescence decreases detectably over several hundred seconds, you have photobleaching. To minimize photobleaching, you must minimize the excitation light by decreasing the excitation slit widths, or by inserting polarizing or neutral density filters. Don’t stop decreasing the excitation light until the signal is stable and doesn’t decrease over several hundred seconds. If you have too little signal now, see (c) below. Remember, you cannot simply correct for photobleaching mathematically. There are free radicals involved. Very dangerous.

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 too high, check with the manufacturer). Don’t mess with the excitation light at this point unless you are well below the level where photobleaching occurs.

Tips for collecting good data:

1. Induce polymerization by adding 0.1 volumes of 10X KMEI (see above). This buffer gives the optimum polymerization rate (See Drenkenham and Pollard, 1984 J. Biol. Chem. for details) and also approximates intracellular ionic composition. Bring all of your stock solutions together and triturate several times for good mixing. Triturating into the quartz cuvette is best. 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. Otherwise prewarm an aliquot of the protein immediately before each assay.

4. Keep track of the dead time. If you are mixing the reagents by hand it will take a finite time to transfer them to the instrument and begin collecting data. Many fluorimeters also do dark count corrections for each data set that take several seconds. To collect analyzable kinetic 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 quartz cuvette to one side and put your pipette tip in the far 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 are bad news.

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 a cuvette washer and don’t spare the water. Rinse with water, ethanol, and then 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. You just can’t.

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 variations in the cuvette position.

10. Make your mind right. Remember what Henry V says: “All else be so, if our minds be right.”


See also:

In Vitro Actin Assembly Assays
and Purification From Acanthamoeba

J. Bradley Zuchero

Methods in Molecular Biology, vol. 370: Adhesion Protein Protocols, Second Edition. Edited by: A. S. Coutts Humana Press Inc., Totowa, NJ