Do you have diatom's mucilage in your samples? 

In my samples diatoms chains  were separated by centrifugation with a previous organic digestion. 

Hi José, thank you for taking an interest in this question.  Yes, my samples are often inundated with mucilage and I would be very appreciative if you could provide additional information on the digestion and centrifugation protocols, if available, otherwise thank you nonetheless, this will put me on the right path to finding a solution!  

The method based on the use of image analysis avoids the problem of estimating material other than zooplankton (Alcaraz et al, 2003. Estimating zooplankton biomass through image analysis. Marine Biology 143, 307-315.).

This website has more information:

 http://www.obs-vlfr.fr/LOV/ZooPart/ZooScan/

Hi Joanna, thank you for sharing the article reference, I will definitely visit this website also.  Much appreciated!

In some cases I have found it convenient to filter subsamples through suitable filter. 

Zooscan is nice but the equipment is expensive.  An alternative is to scan or make a silhouette photograph of the sample and then use WHOI's free Digitizer software to help analyze it. The caveat is that you need access to Matlab to run the program.

This is a slide show describing the method: http://www.whoi.edu/science/B/bioacoustics/SilhouetteDigitizing_files/v3_document.htm

Software and manual download: http://globec.whoi.edu/software/digi_prog/WHOI_Silhouette_DIGITIZER.htm

Kalevi, Nancy, I thank you both for advice given.  Nancy, I've just flipped through your slideshow - excellent!  Some of the screen-shots remind me of analytical techniques employed by my colleagues in our MPA research unit.  One aspect of their research involves studying invert communities from subtidal reefs using images taken with a drop camera.  Thanks again, I am keen to explore this option.

My earlier anwer escaped my tablet prematurely.

I have found it convenient to filter subsamples and pick cladocerans and copepods for biomass determination. A few points are important for successfull results. 1) Filter such water volume which makes detection of animals still easy. 2) The best filters, in my experience, are silver ones (47 mm diameter, 3 um pore size, kept on the top of wet paper to retain zooplankton enclosed within a water film). Their grey color and tough and smooth surface facilitate easy picking. The cheapest, and still useful, alternative is paper filter. Plankton net is possible, but its structure makes zooplankton difficult to recognize. 3) You cannot purchase necessary forceps, but you can easily prepare such. Buy the finest watch maker’s forceps and sharpen those by a fine grinding stone (or paper) under a preparation microscope. Any used forceps of that type are generally useless, because their end has been bent. 4) Use a light guide for best contrast under a microscope.

Kalevi, thank you for providing such detailed advice.  Manually picking will certainly be most effective, although not an option for me at this stage - perhaps when my team is larger.  I also found your advice on sharpening watchmaker forceps useful, particularly damaged ones, as I am sure you know one drop on the floor is normally all it takes to damage these forceps beyond repair.

Dear Shaun,

To my point of view, we first need to know if you want the biomass by family or genus  category, or by the entire zooplankton community. I will answer to the most accurate level of biomass estimation: the family/genus level. I suggest also to use the digital imaginig system (probably another system than zooscan that s pretty expensive) but rather a commercial scanner like Epson V700 and plastic tray) and estimate biomass with a software like silhouette digitizer, the software proposed by Joanna Strzelecki or zooImage. The last one is able to accurately recognise semi-automatically mesozooplankton up to the family genus level (see Grosjean et al. 2004) and use conversion factors to estimate biomass.  Its possible to automatically discriminate particles other than zooplankton but you will need to dilute quite a lot your sampling to avoid the overlapping between copepods and chain forming diatoms. So you will need to use sometime the method proposed by Kalevi Salonen. You can also use the method of José Pedro Marín to preprocess and clean your sample..Its quite accurate compare to traditional methods. Time to dominate the entire system and software: 2 months. But u will save time for the next surveys.. Hope it will also help you.

All the best

Pascal

Dear Pascal - Thank you very much for taking time to address my question, I will certainly make use of all the advice given and develop a system that will work for our organisation.  You have given me lots to think about and I am looking forward to getting systems up and running.  Have a good day.

Yes, let us know!!

Hi Shaun, I would suggest that you consider employing a digital imaging system. Some of the se packages are relatively easy to master and may in the long run save you considerable time

Hi William, it certainly seems to be a popular solution and one I intend to explore in more detail.  I am in the process of buying new microscopes with quite exciting imagery hard- software components added.  It should not be too difficult then to motivate for software purchases to go with it!  Are you or your students currently employing any of these methods?  Please let me know if you/they do, would like to come by for a long overdue visit! Take care and thanks for the input.      

Hi Shaun, I have in the past employed this approach during my studies in the Southern Ocean. At this point, I unfortunately do not have any students employing this technique. You are always welcome to pop in for a cup of coffee or indeed a visit

Hi William, thank you for replying.  I will definitely come by to have a chat soon.