Hi Alberto!

I usually filter the macroinvertebrate samples in order to remove the ethanol, then I dry them up in hoven at 105°C until constant weight (i.e. some days, depending on how many organisms do you have). After this I suggest to let the samples cool within a dryer (otherwise they could rehydrate) and then weight them with precision balance. I also suggest to remove the cases of Trichoptera (of course before drying them), so that they do not bias the biomass measure.

I hope this helps!

Thanks a lot Gemma Burgazzi! I am only concerned about the fact that the sample can not be inspected in future times after the treatment, but there is no answer to that when aiming at dry weight.

Again, thanks a lot for your very precious suggestion!

Alberto Scotti yes, this is the main issue about the evaluation of biomass. After drying macroinvertebrates you won't have other chances to check them, so you should be quite shure about the identification you did. However, the biomass is an interesting metric, expecially if you also have the abundance. In this case you can calculate the biomass-abundance ratio and gain information about the average size of the organisms.

If you have time I also suggest to you to follow the procedure proposed by Georg H. Niedrist which offers really precious informations about communities.

Georg H. Niedrist and Gemma Burgazzi thanks to both of you for the inputs! I am going to start the identification, meanwhile I will think about being conservative, or not, in regards to the methods you proposed.

Thanks and...see you soon :)

The most importante think is the temperature used for drying

Collect identfied and counted organisms dry and weigt

I would also recommend the method suggested by Gemma Burgazzi and Georg H. Niedrist as it doesn't require destructive sampling. It does take more time, but provides much more information about the community. There are many sources available for general and specific length-mass regressions, but if you don't have them for the specific taxa or sites, length-mass relationships are usually pretty robust to the coefficients used in the regression (so regressions at a higher level of organization are often okay). Further, the largest source of error is most often abundance/density estimates (Méthot et al. 2012 is one source that analyses the importance of specificity of L-M regressions)Article Macroinvertebrate size-mass relationships: How specific shou...

Dear Jim, have you some more papers in this direction

Jim Junker thanks for the advices and the article. I will evaluate the investments of time needed (I have many samples to check), but I am happy to hear there are also not-destructive way of doing it!

And its important to remember that once inverts and other animals are stored in ETOH, they lose some of their solutes, making it difficult to obtain an accurate estimate of biomass by drying and weighing.

To estimate invertebrate biomass, I measure the length of the first 20 individuals of each taxon per sample. Arthur Benke et al. 1999 (Journal of the North American Benthological Society) has published length-mass regressions that you can use to estimate biomass from total length. See the appendices.

Lusha M Tronstad thanks for your input! So, you measure only 20 individuals as a proxy, but then you also take into account the total abundances, I guess?

Seconding, Lusha M Tronstad 's suggestion. For samples with large numbers of individuals, I measure the first 20-30 individuals (assuming random picking) and apply the relative size distribution the the total taxon abundance. Also, you can cut processing time by separating samples into sizes by filtering through different size sieves. With the above, you can also sub-sample especially large samples (1/2, 1/4, etc.) and scale up. I often separate samples into size fractions (>1mm and

Hi Alberto, I measure the first 20 individuals I encounter (randomly) and use that as a sub-sample. From those individuals, I calculate the mean size of the taxon and calculate biomass based on those published length-mass regressions. Once you have calculated a mean individual biomass, you can multiply mean individual biomass (mg/individual) by total density (individuals/m2) of the taxon to get biomass (mg/m2) of the taxon. I hope that is helpful.

Lusha M Tronstad and Jim Junker everything clear! Nice suggestions and debate, thanks to everybody.

See here for an example of the length-weight technique, applied in the example to zooplankton but applicable pending the level of detail you desire for your estimate:

Article Length-Dry Weight Relationships of Some Freshwater Zooplankton

1. If you want to estimate biomass, regression equations are a good way to go. There are a number of equation sets out there: Benke et al 1999 and Burgherr and Meyer 1997 are good broad scale data sets. When using equations be sure to use the appropriate value for the intercept (typically denoted a). Sometimes ln(a) is reported as "a" - ln(a) will be -4.0 or so and a will be 1.0x10-6 or so. They will get you to the same number in the end, but you need to be sure you use the right equation form. I would be hesitant to try and develop your own L-W regressions from those samples for the reason provided by Scott Tiegs.

2. I would still advise getting some dry weights so you can validate that the regression equations you are using are providing a 'reasonable' estimate. The masses will be off since they are preserved in ethanol, but they won't be wildly off. An L-W equation that under-predicts the mass of preserved specimens by 50% is just as big of a problem as a poor estimate of density when you go to scale up to the community.

3. Propagate your error / uncertainty through the entire process - don't ignore the error terms in the published regression equations. Apart from the regression equations, you have uncertainty in multiple measurements that need to be integrated / propagated.

4. For getting the mass of the community, if you have many samples/sites (hundreds) to get through you, can use the approach suggested by Jim Junker . However, this can result in greater uncertainty about your estimate depending on the taxonomic level to which you are going and whether or not the taxa are semivoltine or multivoltine - if you have mostly univoltine species this is less of a concern.

5. If you have less than 100 sites / samples, to reduce uncertainty, I would suggest directly measuring all animals that you identify (unless they are all univoltine, in which case Lusha M Tronstad 's method is most efficient). A simple solution for measuring the length of all animals is to take a picture once they are taxonomically sorted. You can then use ImageJ to quickly measure lengths with high accuracy. If your dissecting scope is equipped with a camera, this is simple and you are probably already doing it. If you don't have that setup, you can use a flatbed scanner and put the animals into a container with a thin flat transparent bottom. I use transparency film glued to a frame. A 1200 DPI optical resolution scanner will give you more resolution than you could reasonably need (even for daphnia). A good quality used scanner can be found for less than $10 in the US.

6. Be sure to remove trichoptera from cases before measuring.

Article Length-Mass Relationships for Freshwater Macroinvertebrates ...

Article Regression analysis of linear body dimensions vs. Dry mass i...

Also if applying equations across spatial gradients or from different parts of the world consider:

Article The biogeography of insects' length–dry mass relationships

Benke, A. C., A. D. Huryn, L. A. Smock & J. B Wallace. 1999. Length-mass relationships for freshwater macroinvertebrates in North America with particular reference to the southeastern United States. Journal of the North American Benthological Society 18 (3): 308-343. DOI:10.2307/1468447

Becker, B., M. S. Moretti, & M. Callisto. 2009. Length-dry mass relationships for a typical shredder in Brazilian streams (Trichoptera: Calamoceratidae). Aquatic Insects 31 (3): 227-234. DOI:10.108 0/01650420902787549

Burgherr, P. & E. I. Meyer. 1997. Regression analysis of linear body dimensions vs. dry mass in stream macroinvertebrates. Archiv fur Hydrobiologie 139: 101-112.

Thomas B Parr thank you for your exhaustive summary!

In Finland, we use wet weights e.g. for profundal macroinvertebrates of lakes. This is a one option. First, you could sort animals to taxonomic groups wanted. Second, you move animals to be weighted from alcohol into water. Third, before weighting them with scale you should dip animals in some kind of paper.

Another option would be to measure lengths of all individuals and use length-weight relationships from literature to convert dry weights.

First method (wet weight) is less time demanding, but more inaccurate. Dry weight method is more time demanding and more accurate.

Significant summary nevertheless the final decision is dependant of money afforded with consequences on equipment and man powe salaries