Biology Reference
In-Depth Information
optical sections, thereby excluding the out-of-focus information. TIRF microscopy is
also an option for plant biologists needing optical sectioning. Chlorophyll is the main
pigment that autofluoresces red from
550 to 750 nm when excited with either the
standard 488 or 543-nm laser lines. Band pass filters or acousto-optical tunable filters
are required to remove the autofluorescence when imaging mCherry.
Each experiment requires different microscope and camera/PMT settings that
must be determined prior to data collection. Many confocal microscope software
packages have generic settings for each laser and/or probe that are helpful starting
points. The magnification and imaging interval (single time point or time lapsed)
should be determined first. When imaging, the usual goal is to collect as much signal
over background noise as possible, while using the least amount of laser light possible
in order to avoid bleaching the fluorescent protein or stressing the living specimen.
Whenmaximizing signal to noise on confocal microscopes, the PMT settings and laser
intensity are linked. Increasing the PMT gain amplifies all signals, including the back-
ground noise. Frame and line averaging can be used to reduce the background noise.
Increasing the laser intensity usually generates a higher signal-to-noise ratio, but
bleaches the probemore quickly. The amount of acceptable bleaching is often dictated
by the imaging interval. Probe bleaching is not a concernwhen imaging at a single time
point and so laser intensity can be increased, rather than PMT gain. Photobleaching
and plant health must be considered when collecting a time-lapsed image series.
One must usually sacrifice some image contrast by decreasing the laser intensity
and increasing the gain to image over time. It is difficult to image fluorescent reporter
fusions that are not abundant. One strategy to image dim reporters is to sumor average
images taken over time. The specimen is likely to drift while imaging, and this can be
corrected after image acquisition with software such as the StackReg plugin for Ima-
geJ (
http://bigwww.epfl.ch/thevenaz/stackreg/
)
. Deconvolution software can also be
helpful to remove some of the background fluorescence from the data sets.
To measure the dynamics of microtubules and MAPs, we routinely used a 63
water immersion objective and electronically zoomed 2-3 times so that a single
light-grown hypocotyl cell filled the field of view. When imaging microtubules
and MAPs, we have found it useful to measure the signal intensity using a line scan
tool. To start, draw a line across the entire cell and quantify the fluorescence. Next,
adjust the PMT gain so that almost no pixels are saturated, and ensure that the base-
line is slightly above zero by adjusting the offset. Now measure the signal of a few
single and bundled polymers by drawing a line across several microtubule structures.
To ensure a wide and linear dynamic range, an incremental stepwise increase in sig-
nal should be observed due to the bundled microtubules (
Eisinger, Kirik, Lewis,
Ehrhardt, & Briggs, 2012
). A linear increase in signal is especially important when
quantifying fluorescent signal over time, such as in fluorescence recovery after
photobleaching experiments. However, because some microtubule bundles are com-
posed of numerous microtubules, we have often found it necessary to saturate the
fluorescent signal of higher order bundles in order to distinguish single microtubules.
To ensure that sufficient signal over noise is collected, it is important that the pixel
value of single microtubules is at least 2-3 times the background value.
