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As far as what confocal to get, it is hard to give you any specific advice. It is 
like buying a car. It depends what your users need.
I like them all, each of the big four has their strong points and offers a range 
of models. Some of the lower-end models are upgradeable, some are not so 
easily. And, as was mentioned in this thread, there are other options, like the 
Thor Labs kit, and few others if you are looking for a single-point scanning 
confocal system. Spinning disk/micromirror instruments are yet another group. 

 
We have a dual approach for training.

There is a graduate - level course where students learn the theory of image 
formation and a range of light microscopy techniques, and receive practical 
training on a standard light microscope (BF, DIC, phase contrast, 
fluorescence) as well as on a confocal microscope.

For users that just want to learn how to use the confocal microscope I do the 
training (2 people max) in two sessions, ~2 hours each.
We start with general and lab safety, biosafety rules (we are BL-1), each new 
user is required by our university to have passed an online laser safety 
training. 

I have a short guide for users to download, it also includes startup/shutdown 
procedures.

In the first session we cover the operation of the microscope, what do those 
buttons do, setting up Kohler illumination, using standard epifluorescence to 
check the sample, and basic functions in the software: scan size, speed, 
setting the detector voltage and offset for maximum contrast and to avoid 
saturation, choosing fluorescent dyes from the dye database, confocal zoom 
and setting the pixel size for Nyquist sampling, sequential scanning to minimize 
spectral cross-talk in multi-labeled samples. We also cover how to deal with 
weak signals and noise by changing scanning speed, Kalman filtering, opening 
the pinhole.
Usually in the first session we only use the 10x and 20x dry objectives and a 
standard teaching sample (I use pollen grains).
We also cover the file formats, the importance of saving the original data files, 
adding a scale bar, exporting the images for a Powerpoint resentation.

I like to demonstrate the effect of the DIC prism on the confocal image 
resolution and signal brightness. 


In the second session we cover the use of objectives with coverglass 
thickness correction collar (dry 40x and water immersion 60x), using oil 
immersion objectives, and general rules how to clean up after using the 
immersion optics. Z-stacks - setting the z-step size to Nyquist, adjusting the 
laser power or detector voltage to compensate for loss of signal  with depth.
Time lapse imaging.
Towards the end of the second session user get to look at their own 
specimen, if they have one, and get help with optimizing the imaging 
parameters for their sample. 

What exactly is covered depends on each user's need. Some people will never 
use a water immersion objective, some people only look at thick live samples  
and will not use the oil immersion lenses. 

I inform the users about the more advanced features, like the photon counting 
mode, bi-directional scanning, line-scan and point-scan, FRAP, FRET and 
acceptor photobleaching, using a SIM scanner (it is an Olympus FV1000) for 
photobleaching/photoactivavtion, Multi-area Time lapse (uses the motorized 
stage), assembling complex imaging sequences ("macro"), Raster Image 
Correlation Spectroscopy, ... but for 99% of the users this would be more than 
they need at the moment and it does not make sense to train them for this. 
     
After the two training sessions the users should be able to use the confocal 
microscope more or less independently, with some occasional help. They can 
use the instrument during normal operating hours (~9am-5PM).  After one or 
more sessions, when they are comfortable in using the microscope and do not 
need any more help, they let me know and we go through a short practical 
test, where they show me they can indeed do the imaging without 
endangering themselves or the microscope.  After the successful practical test 
they become fully qualified users, they get access to the online scheduler and 
get a key to the microscope, so they can use it any time.

Even after the training is completed I still like to check on my users, ask how it 
goes if they have any issues with their images, and dispense some unsolicited 
advice, e.g. that using Alexa488 and Cy3 for dual labeling and colocalization 
analysis is not the best idea, there are better dye combinations.
 
Overall this training scheme works well, we have had minimal problem with 
users doing bad things to our microscope. Sometimes I train the students and 
then they realize that they use the microscope so infrequently that they are 
better off just letting me do the imaging once or twice a year, rather than 
trying to re-learn what they have forgotten since the training.
 
With Regards,


Stan Vitha

Microscopy and Imaging Center
Texas A&M University
http://microscopy.tamu.edu

On Mon, 18 Mar 2013 14:56:50 -0700, jmkrupp jmkrupp <[log in to unmask]> 
wrote:

>*****
>To join, leave or search the confocal microscopy listserv, go to:
>http://lists.umn.edu/cgi-bin/wa?A0=confocalmicroscopy
>*****
>
>Greetings
>
>Just a quick note to ask if anyone would like to give me some advice about
>getting started with a confocal training program.
>
>We do EM and LM already, thinking of adding LSCM. Any advice about must
>includes,  good practice specimens , techniques etc?
>
>How about insights into instrurments, how basic can we go?, what cost range
>should we be thinking.
>
>Thanks
>
>Jon