Hi Rebecka

as you can see this is a subject that has tortured scientists working on filamentous fungi for many years.

The suggestions above are all good. In my experience dry weight works best then wet weight (eg pressed in Miracloth), packed volume works if your fungus will spin down easily - they don't all do this, DNA is OK (RT-PCR of a genomic target) and others have used ergosterol (OK if you have ready access to analytical HPLC). Finally OD600 can be used but only on thin well dispersed cultures - ie no lumps!

However it looks from your post that you may have another problem. If you are homogenising then aliquoting equal amounts into different tubes but are still gettting differences then you need to carefully consider the homogenisation technique. Also pipetting homogenised fungi can be problematic - the lumpiness issue again. A few tips on this might be to cut the ends off gilson tips or always use a wide bore pipette. Others weigh wet mycelium to quantify the amount then cut equal weights from the mass of mycelium, resuspend by vortexing and inoculate with this. This is easier than it sounds and allows for some reproducibility between experiments - it works well for us for RNAseq replicates which is fairly demanding.

Good luck!

Hi,

for the OD you have to validate the method to convert the OD number to number of spores or CFU...

In my previous years, when I was working with fungi, I counted the number of fungal conidia per mL of solution with counting chamber under microscope. I let the fungus grow on agar, then scrabbed the mycelia with conidia, filtrated it and took subsamples, put them into counting chamber and microscopically counted...

It is a very interesting and hot topic with filamentous fungi cultivation... First, it would be useful to know if your fungus produce spores. In that case, usually different scientists use this state to prepare standard inocula, e.g. 10^8 spores/mL in the inoculation suspension. I frequently use a 10% v/v inoculum. But in my case, the fungus is not able to produce spores, only mycelium. Then, I tried the homogenization protocol with a common hand blender. Probably a technique similar to yours. This protocol works quite well. At first glance, I would not recommend you to take OD measurements. I tried, and it did not work properly for standardization. I suggest you to prepare agar plates from this fungus, that should be grown in a culture medium, and for a given time, always under the same scheme (e.g. Czapek, 48 h, 30 degrees). Then, try to take a standard number of mycelium-covered agar plugs (taken with a cork borer or a sterile needle) of a certain diameter.

Another option is dry weight, which can be quite precise, but the problem is that it takes certain time (e.g. at least 5-8 h) to get the constant dry weight... This works especially if your fungus does not produce polysaccharides or other metabolites attached to mycelium... of course. And finally, there are indirect techniques which might also work: glucosamine determination, or ergosterol quantification, but we tried them for solid state fermentations, when there were no other ways to solve biomass determination...

Cross your fingers and try this simple tips. They might work for U also! Best wishes ([email protected]; [email protected])

Normally we used blood cell counting chamber to count the cells of fungi or Collect mycelia by vacuum filtration and the same weight of mycelia were weighted for further study.

Filamentous fungi grown on agar plates form spores. Saline/tween solution (10ml) is pipetted on the spore-mat and spores are scraped off using a bend inoculation needle. The spore suspension is transferred to a sterile bottle (stock). The concentration of spores in the stock is determined by counting the sample diluted with saline/tween solution in a haemocytometer. The concentration of spores is calculated depending on the haemocytometer (depth and width) used. Approximately, 1000000 spores can be inoculated into 5 mL pre-culture complete medium, which will be transferred into the main culture (200mL) after overnight incubation. The number of the spores can be adjusted according to the fungus used.

Hi,

From my experience the neubauer method is a good approach if you are looking for fungal conidia,

(http://www.ruf.rice.edu/~bioslabs/methods/microscopy/cellcounting.html)

However, from your description, you are using mycelium, so even if you 'homogenize' it, it will still be carrying lots of different cell sizes and cell agglomerates, which makes this technique a bit less usable, furthermore, you might have a mix of conidia/mycelium cells.

Regarding the OD, i also thought of that at the time, but some fungi (like A. niger) produce lots of pigments that will eventually change the OD of your samples, so OD usability will also depend on the organism you are using...

What i did use, with good results was flow cytometry, since you can count and measure the number of cells per given volume of sample, and if you do it properly (by filtering your counts in the cytometer according to cell size and shape) you can actually have a very robust cell count ! You can check my paper here :

http://www.sciencedirect.com/science/article/pii/S0964830512001084

In case you need further help, just let me know =)

OD is the best method, but the growth rate can be estimated even from the metabolite secreted by fungus

thanks all of you for your answers! I will have to think oabout them for a little while and then i might get back to you.

OD would be the easier for you and simpler.

The usual way is measuring dry cell weight. (Filtrate standard amount of sample from the medium, let it dry, than measure.)

There are some good pointers in all these answers here. My first question: what filamentous fungi? Not all fungi sporulate well. Even if they sporulate, the hydrophobicity of the surface layers may cause them to clump together in any aqueous buffer suspension making OD measurements or hemocytometer counting difficult. A general rule of thumb is to collect the spores in a buffer containing a surfactant, such as 0.05-0.1% Tween-20. It helps in keeping the spore dispersed. The spores can then be counted under the hemocytometer. The OD measurement would vary due to multiple factors, such as type of fungus, size of spores etc., and it doesn't always correlate well with a viable count. For some fungi whose genomics are relatively well-defined, the newest method is to estimate the count from the total quantity of some marker.

I read this on another question and might be useful to you:

BY:Sanjay Antony-Babu · 14.36 · INRA - Institut National de la Recherche Agronomique

1. Oatmeal and cornmeal agars enhance spore production in most filamentous organisms, both fungi and bacteria.

2. You can place sterile filter papers on agar plates and inoculate on top of the paper. After growth you can slowly peel the paper off the media picking off the spores.

3. You can pour Tween 80 or 0.2% SDS (this reduced hydrophobicity of spores) on fungal colonies, make a suspension and filter it through sterile cotton wool. Cotton wool traps hyphae which are long and lets circular spores to pass through.

4. If you need to remove spores and be quantitatively precise - remove a block of fungal colony, place it in 0.2% SDS solution (in a falcon tube) and shake it overnight. This will slowly release the spores which you can quantify using a haemocytometer.

I have used all these methods in the following papers:

Antony-Babu & Singleton (2009) Effect of ozone on spore germination, spore production and biomass production in two Aspergillus species. AVL International Journal of General and Molecular Microbiology 96(4), 413-422

Antony-Babu & Singleton (2011) Effects of ozone exposure on the xerophilic fungus, Eurotium amstelodami IS-SAB-01, isolated from naan bread. International journal of food microbiology 144 (3), 331-336

Good luck!

I forgot to mention, another advantage of using a flow cytometer is that you can actually run the your liquid medium directly through it (if fluid enough of course). If you use simple cytometer mesh filter, with the mesh-size of hyphae cells and/or spores, you can actually detect them and count them for a given volume of medium on the cytometer, and filter out cell types according to different parameters!

I think it's wonderful that there are so many people out there wanting to help! :) I think I might have to be a bit more specific with what we want to do. We stury Postia placenta, a brown rot basidiomycete, which is a filamentous fungi that doesn't sporulate very easily. We study what this fungus doesto wood, which means we have wood pieces on a little bit of soil in petri dishes and then we add the fungus. You could of course out an agar plugg on top of the wood samples, but i dont think that's very easy to standardize eather. So instead we put agarplugs in a liquid medium (malt) and let them grow for two-three weeks. They form a big lump of mycelia floating on top of the medium. When it was time to start the experiment we used a hand mixer to try to homogenize the fungus in the medium, but it was really difficult. When we decided to stop there were still some bigger pieces, but those we didn't use of course, but the small, pipettable pieces of course also differed markedly in size. Then we added 1 ml medium to each wood piece. At the time it felt like we distributed the medium quite equally, but we can see now when we harvest our samples that we have a lot more growth on certain samples than on others and the wood pieces are also degraded to different extents. We have two samples in each petri dish and it can differ quite a lot between the samples in the same dish. This means we need to have a method that is rather quick and easy, and we don't have aflow cytometer. Dry weight right now seems like a good alternative. Erzsébet Sándor, how would you dry the cells? For how long?

Does nyone have a good idea for how to get rid of the bigger pieces? Is there any kind of filter or so that might be useful?

Thanks again everybbody for wanting to help!

Dear Ms. Ringman,

May I suggest that you read the following paper

J Clin Microbiol. 2001 April; 39(4): 1345–1347.

doi: 10.1128/JCM.39.4.1345-1347.2001PMCID: PMC87937Inoculum Standardization for Antifungal Susceptibility Testing of Filamentous Fungi Pathogenic for Humans

In this paper it describes some means to measure numbers of filamentous fungi and discusses the accuracy of different methods.

I hope that this is useful to you.

Best wishes

Thank yoo so very much, Gerard! I will check this out.

I would like to know the reason why you are using liquid culture. As an inoculum?

Philippe, since we are doing this on soil we need to put the inoculum on top of the wood sample and using liquid cultures has two advantages, one is that the fungi will be as evenly spread on top of the wood sample as possible and two is that the soil takes a lot of space and there is no room left for an agar plug on top of the wood sample... Those were the original reasons, but after some thinking we also feel that using liquid cultures would make it possible to make sure that we add the same amount of cells to each sample too - if we only could find a way to measure the concentration. Even if you grow your agar plate the same number of days and make all pluggs the same size, that doesn't make sure all have an equal number of cells....

Rebecka, I understand the pb, you need a standardized inoculum. I guess that your fungi does not sporulate easily. Otherwise, it is very easy to get a standardized spore suspension. Is your fungus sporulating on solid medium?

Our fungus is unfortunately not easy to sporulate.

I agree with Gerard Colleran. In my experience with antifungal susceptibility, at the point where we needed to adjust our inoculum for the assays, we would prepare a lot of cultures and the scrape the mycelia with a pasteur pippete and a little bit of sterile saline solution. We would pour the scraped material inside a sterile glass syringe that contained some gause in order to filter mycelium and let spores through. We would then count with an hemocytometer as well as viable count in agar. Both quantities should be similar.

I hope this helps.

:)

Flow cytometery is the best option

I've had the same problem with non-sporulating species. We tried OD as well with inconsistent results. What worked better for us was grinding the sample from our primary culture in liquid nitrogen and then lyophilizing it. We then used a fixed weight of the lyophilized sample as the inoculant. It is a bit more work but on the positive side you can prepare a large batch and store it easily over the course of several experiments and your inoculant remains the same throughout.

Dear Rebeka,

Regarding the filter paper for fungal dry cell weight, I can not find the type right now (sorry). Try the ones, with pore size bigger than for bacterial and sterile filter, and the filtration is rapid enough. If you are in a hurry, try MYRACLOTH (it is a pretty expensive, gut the filtration is extremely quick).

Cell amount measurements, like flow cytometry and turbidometry are developed for unicellular organisms, and not suitable for filamentous fungi.

All the best for your work!

Erzsebet Sandor

OD is very problematic with filamentous fungi. Flow ctometry would only work on small germlings and you would likely need spores for that. When I have worked with species that don't sporulate, we would often propagate them from agar plugs using a blender. Take a small colony and grind it in a tissue grinder in liquid media. This will cause the culture to grow from many small points instead of one giant ball. You could collect this and use it for inoculation in lieu of spores.

Another method to measure fungal content though that no one has mentioned is to measure ergosterol content. Ergosterol is essentially the fungal version of cholesterol and therefore you can correlate fungal content with ergosterol content in a mixed sample. Here is a reference:

http://www.apsnet.org/publications/phytopathology/backissues/Documents/1979Articles/Phyto69n11_1202.PDF

If, after trying all the tricks to induce sporulation (nutrient deficient media, etc), you cannot get your bug to make spores, then the procedure that Corinne suggested would work well and give you the most reproducible method for estimating inoculum.

Packed cell volume (PCV)could be suitable tool to measure the fungal contents as it does not take more time to determine the contents. You can also correlate the PCV to dry cell weight of fungal biomass per unit volume.

Dear Friend, I recommend an easy methods for the evaluation, it is using a Neubauer Chamber or Homocytometer that already used to evaluate red cell of human blood. please try it.

Dear Rebecka please see attached file that show using Homocytometer step by step

Yes Majid Azizi:s answer is only suitable for unicellular. microorganisms not filamentous fungi The suitable answer may be either dry biomass or go for PCC.

Pl Read it as PCV ( Packed Cell Volume).

It is always more complicated with filamentous fungi that don't sporulate. (I worked both with Botrytis and Sclerotinia = many vs no spores).I like the idea of Kenneth and Corinne to make a primary culture with many small fragments. This will give a more even growth in your bigger culture. This also helps becasue you can grow it shorter and all cells are closer in age, the cells in the centre of your big ball of mycelial aggregates may be much older or nearly dead already, so even when cut to the same size they will grow out differently. Let us know which method worked!

Hi Rebecka

as you can see this is a subject that has tortured scientists working on filamentous fungi for many years.

The suggestions above are all good. In my experience dry weight works best then wet weight (eg pressed in Miracloth), packed volume works if your fungus will spin down easily - they don't all do this, DNA is OK (RT-PCR of a genomic target) and others have used ergosterol (OK if you have ready access to analytical HPLC). Finally OD600 can be used but only on thin well dispersed cultures - ie no lumps!

However it looks from your post that you may have another problem. If you are homogenising then aliquoting equal amounts into different tubes but are still gettting differences then you need to carefully consider the homogenisation technique. Also pipetting homogenised fungi can be problematic - the lumpiness issue again. A few tips on this might be to cut the ends off gilson tips or always use a wide bore pipette. Others weigh wet mycelium to quantify the amount then cut equal weights from the mass of mycelium, resuspend by vortexing and inoculate with this. This is easier than it sounds and allows for some reproducibility between experiments - it works well for us for RNAseq replicates which is fairly demanding.

Good luck!

It's really nice to see that so many people still are trying to solve this problem for us! :) Thank you so much all of you!

The most appopiated method is the filtration to masure dry weignt, and the mycelia package, fixing volume sample, centrifge speed and time.

I always use the Neubauer chambler to count the fungi. It's easy and fast and you have reproducibility in your experiments. I used also dry weight, but I use it in the end of a culture, for example. To quantify the fungi, before starting an experiment, I always use the Neubauer chambler. I am sending you a protocol, if you want.

To inoculate fermentations you must not homogenizate the inoculoum. You should make an inoculum for aech fermentation. Tje homogenization could destroy your micelium. Not all the fermentations broth produce enough spore number. The inoculation of fermentacion can be done by harvesting spores from solid cultures. You can have normalized spore suspentions using the Neubaer Chamber. Then, you can use this spore suspention to inoculate your fermentation broth. For inoculum you must to adjust the fermentation time for the inoculum and the inoculate the fermentation broth without homogenizating the inoculum. Using the same fermentation time for inoculum almost always you have the same biomasa.

If you can grow the fungi on a solid medium and it will grow outward in a circular pattern; try taking small plugs from the leading edge of a the growing mycliea.

Usually we'll use a sterile inverted 10uL pipette tip to make a 5mm diameter agar+myclia plug.

Thanks Bryan for such a simple suggestion! :) the more we think about it the more we realise that the variations in the wood samples that we inoculate with the fungi are bigger than the the variations in the amount of fungi we inoculate... So this way may be good enough!

Since there is no ready method for working out the quantity of propagules in an aliquot of homogenized filamentous fungus, your best bet is to ensure that for each experiment, you use a highly standardized method for the production of your inoculum i.e. start with a standard amount of propagules to start your first liquid culture (this may be a set number of plugs from the edge of growing colonies; plugs of standard size each time); set your first liquid culture to about 20 mL in a standardized flask size and type e.g. 100 mL Erlenmeyer with a standardized closure e.g. Sigma silicon foam closure. After a set incubation period of fixed length (keep it the same each time), homogenize the contents and inoculate into the next sized flask culture (may be now to 50 mL in a minimum 250 mL sized flask) at a rate of 10% (v/v). Then incubate a second time. You then homogenize that last batch and use it for your experiment. The objective here is to passage through sufficient number of flask cultures to even out differences. Having three replicates at each level of culture helps.

I presume that you fungus does not sporulate but if it does there are ways to standardize aliquots of spores. Check out the methodology in:

http://clemkuek.com/papers/kuek&arm.pdf

Clem

Thanks Clem! I think it is amazing how this question has been kept alive for so long now! But this is an ongoing problem for us so we are very grateful for all the tips and ideas we get! I think this method is the closest to what we currently do, but with some refinements. As I've written somewhere above, thebiggest issue is not to get the same amount of fungi each time but to grow them on wood pieces that don't differ too much from each other. So an easy method that will eliminate the greatest issues with the fungi I think is suffcient, like yours! Thanks again! :)

I work with Neurospora and I find that if you are working with conidia, you can just count using a haemocytometer and viability can de determined using methylene blue. I usually get about 100 million conidia/slant when grown on Vogels minimal media. I find that body mass is the only accurate measurement for mycelia but you may also try a vital dye like Alamar blue to determine relative cell viability. Vital dyes may give false numbers and should be reproduced to trust the results.

Dear Rebecka + All,

Great day.

I had tried to grow the Rhizopus oryzae in the pellet form using various media.

I had followed a few papers but, it seem not successful.

The media that used were (autoclave before innoculation)

1. PDB (liquid Potatoes + glucose)

2. glucose,

3. glucose + CaCO3,

3. Glucose +CaCO3 + peptone (meat)

pH was adjusted to 5-5.1 using Acetic acid.

conditions: 180rpm

Temp. 37C.

Innnoculum: 5-8ml sterile dH2O washed the 3-5days Rhizopus agar plate (incubate at 37C) to get spore suspension.

innoculumm added into shake flask was 2%, 4, 6, 8, and 10%.

Shake flask working volume was 50ml

However, most tested media produced clump cell, growing in pellet cell was considered less growth...

Based on your experience....

1. Is the temperature too high?

2. Rpm to slow/fast?

3.pH?

4. Innoculum should further dilute at least 10x, 100x?

Thank you for sharing ur experience here.

Jacky

Normally yeast can counted as individual cells. But usually dry/wet weight alone used in mycology research.

I have experience growing white rot fungi to rubber wood block (6cmx6cmx6cm). Put 2 block in 1 plastic bag and poured about 150 ml of malt extract broth into the plastic bag and tight with rubber band. Autoclave at 121 C for 30 minutes. after cooling overnight, inoculate with mycelium mat from agar. Incubate at 25C for 2 month and will get the fungus colonise into rubber wood block.

Do you mean "solution" of filamentous fungi? Is it not a suspension? Pedant I know...

You can use a solid medium such as PDA, after incubation you can remove the fungus from the surface agar using a PBS solution. Usually I mix very well the suspension before doing my experiments, you can use ultrasounds or a stirrer such as an ultra-Turax.

Russell: Can you explain why are you pedant?

I meant that the authors just used a the wrong term viz suspension and solution. And I was being a little pedantic to point it out.

Ok. A good point. We are talking about solid matter into a liquid phase, not ions or molecules. A suspension not a solution. But, why pedant? You are just right.

Copy that...

A little more... The original question was: How can I measure the amount of cells in a solution with filamentous fungi? The sentence points "with" not "of". I understood that the solution was the culture medium not including the fungus. But reading the explanation of Rebecka ("...use a mixer to try to homogenize the solution...") she makes some mistake. I think that it is the result of the horrible grammatical style that contaminate e-mails, facebook, RG, etc. But, who is without sin be the first to throw a stone... However your observation is very good! because I think this is not a site just for "I ask you answer". The main richness of RG is, probably, teaching. There is too many young people that are excited to learn. In this sense the comment you meant about precision in scientific language is very valuable! Cheers.

As an alternative, glucan amount can also be tried to determine inoculum amount in the medium. But the medium composition is very important to use this method. Some media can contribute to determined glucan amount.

Dear Rebecka,

I'm I understood correctly? Do you want to inoculate a liquid growing media with a spore suspension? If it is so, you can use the haemacytometer to count the number of spores.

Dear Ana-Christina,

No, since our fungus doesn't sporulate easily we need to inoculate with mycelia and we're having doubts whether we homogenise it well enough and are also a bit concerned that we might not be able to add wequal amounts from experimetn to experiment.

Again I'm amazed by the number of people commenting on my question, which I posted about a year ago now. I thank you very much, all of you, for taking your time to try and help us out. As I've said earlier, it has turned out that the difference in amount of fungus in our SUSPENSION (;)) is not as important as the differences in the wooden blocks we inoculate. Therefore we keep on as ususal witht the fungal suspension. But all tips and trick are gratefully excepted anyway! :)

I agreed with Rebecka replied.

So far, any good method you have found to perform fungus research particularly the equality amount ?

Hi. Looks like this is already an old question, but for anyone still looking for a solution ... I would suggest that you try adding 0.2 -0.4% agar to your pre-culture medium. (Make sure that you shake the flask while it cools to prevent the agar from solidifying at the bottom.) This provides what appears as a lumpy medium, but usually liquifies nicely once you inoculate it with the fungus. Inoculate with your homogenised mycelia. (Do not homogenise for too long or the heat will kill them, increasing variability in the suspension.) The agar should help keep the fragments dispersed, enabling you the fungus to grow as smaller pellets or even filamentously. This suspension can then be pipetted without further homogenisation (still a good idea to clip the tips or use wide-bore, though). Your mycelia should be in a healthier, more homogenous state and thus replicate aliquots from the culture are more reproducible. Works with many, but not necessarily all fungi. If the agar interferes with your inoculation procedure, this method can also be used to establish pre-pre-cultures, which are used to incoulate medium without agar (e.g. 1:10 ratio), which dilutes the agar in the original pre-culture. The second culture will probably form pellets, but typically smaller and more uniform than when inoculated with homogenised mycelia directly. The agar is inconvenient if you need to filter your samples before inoculating. For some strains, gelatin can be used instead of agar. Larger concentrations are needed, but the fungus should break it down as it grows.

Hi, this sounds really interesting! I think we'll have to try this! Thank you!

I am in accord with @Marilyn

Dear Rebecka

I could not understand your question.Do you want to count the amount of spore in your solution or the amount of spore plus mycelia. But if you want add the right amount of spores, you can culture the fungi on solid media and use haemacytometer to count the spore concentration and then by providing the dilution serials can reach to wanted concentration of spores.

Dear Rebecka

your question is very interesting. In the case of fungi with mycelia it is difficult to homogenize as the insoluble mycelial particles do not remain in the homogenized state. Every sample  may contain  a different  amount of mycelia.

I think if you centrifuge the mycelia and separate the packed cell volume and add the known weight of mycelia to each sample might give you some uniformity. You can try if it works.

Please see the approache of the papers below:

Fateixa S, MC. Neves, A Almeida, CP Neto, J Oliveira, T Trindade (2009). Antifungal activity of SiO2/Ag2S nanocomposites against Aspergillus niger. Colloids and Surfaces B: Biointerfaces, 74: 304–308.

Pinto R, Almeida A, Fernandes SCM, Freire CSR Silvestre AJD, CP Neto, Trindade T (2013). Antifungal activity of transparent nanocomposite thin films of pullulan and silver against Aspergillus niger. Colloids and Surfaces B: Biointerfaces, 103:143- 148.

Dear Rebecka,

Unfortunately there is no accurate method for determining the number of cells (probably you mean CFU/ml or CFU/g) because a fungal colony contains mycelium that is very difficicult to split and also conidia and/or spores.

In almost all studies concerned with fungi, fungal conidia or spores are inoculated. Biomass can be evaluated by the diameter or radius of the colony in solid medium, by dry weight in liquid medium.

In my experience working with fungi I have saw the fungi growth as pellets in agitated liquid media. The pellet size depends on the number of spores inoculated broth. A low number of spores produce a loose big pellet. A high number of spores produce a tight small pellet. Then,105 spores/ml as final concentration  in the media culture, used as inoculum, is an appropriated condition to begin the fermentation. I suggest use single inoculum for each bigger culture, adding to each one the same number of spores. It is necessary to agitate very well (200 rpm).

Dear all experts:

Greeting.

I am producing lactic acid by using fungus pellet together with insoluble lignocellulosic biomass via simultaneous saccharification and fermentation (SSF).

Adding calcium carbobnate, CaCO3 after 6 h SSF process.

I found that it was impossible for me to determine pellet biomass as the insoluble substrate, CaCO3 and fungus were mixed together and hardly to separate.

I have tried centrifuge the broth and only ale to separate the supernatant and pellet (fungus + biomass + CaCO3).

Any good idea / method to determine the biomass?

or....

the biomass determination in this case is meaningless since the target product is lactic acid?

Please give your opinions.

Thank you.

Hi. A suggestion for Jacky Lai and the problem of biomass + fungus + CaCO3.

It is good that your results are not dependent on an accurate measurement of fungal biomass, however, it is often useful to understand production in terms of the amount of biomass needed to achieve a particular result, so it is understandable that you would like some measure of fungal biomass, even if your product is lactic acid. Unfortunately my suggestion will take some time. It will not be completely accurate, but will at least provide an estimate. Several variations on this suggestion can also be imagined...

1) Measure total biomass by either filtration (e.g. 5 ml sample on a glass fibre filter) or centrifugation and washing.

2) Take a total sample (fungus + biomass + CaCO3 + liquid) of known volume (e.g. 1 ml) and inactivate the fungus and any enzymes by e.g. heating at 100°C for 10 min.

3) Add crude cellulolytic enzymes (e.g. something like Celluclast with Novozyme 188, plus hemicelluloytic enzymes if needed) and incubate 2-3 days at 45°C, preferably with mild shaking. This should hydrolyse most of the cellulose (and hemicellulose, if present and appropriate enzymes added). Most of what remains should be fungus + CaCO3 + lignin.

4) You know how much CaCO3 you added to the culture. Calculate how much acid (e.g. H2SO4) would be needed to dissolve the CaCO3 and add an appropriate amount of acid to the enzyme treated sample. Mix well.

5) Collect the remaining biomass by filtration or centrifugation (with appropriate washing). This should provide a measure of biomass + lignin. If lignin is not modified or removed during the culture, the lignin contribution should be constant and can be subtracted to estimate the biomass.

6) Before dissolving the CaCO3, you could also remove a sample of supernatant for sugar analysis to estimate the cellulose/hemicellulose degradation and to use as a check with comparison to the total biomass sample.

If your fungus is efficient at degrading the biomass you are cultivating it on, you can also skip the hydrolysis of the cellulose step (and the heat inactivation step) and directly treat the sample with acid to dissolve the CaCO3. Then collect samples for biomass DW as usual (filter or centrifugation with washing). Initially the weight may increase as the fungus grows on easily obtainable sugars, more than it has digested the fibres. As the hydrolysis (SSF) procedes, the biomass should decrease to provide you an estimate of the final biomass once all the substrate has been consumed. If you see an initial increase in biomass the amount of increase should reflect fungal growth and provide an initial biomass estimate. This also is affected by the relative amounts of lignin, which should be determined for fungus-free samples.

Note that the final measured biomass is only an estimate. Both the heating step and also the treatment with acid may cause cell lysis and loss of some cellular components which are then not included in the final biomass measurement. Alternatively, you could consider indirect biomass measurements, but these also are estimates.

I hope this inspires you to find other solutions also.

Kind regards,

Marilyn

Hello i want to prepare spore suspension from cylindrocladium parasiticum from a 14 day old v8 culture , my questiions are;1. how do i prepare the suspension

                                                                     2. Can i inoculate directly without determining the spore concentration.   

Hello Rebecka Ringman

We can obtain A conidial suspension by scraping mycelia and spores from plates of actively growing fungal cultures into sterilized water and filtering the suspension through four layers of cheese cloth to remove most of the mycelia. Then filter the spore suspension,  centrifuge at 2000 x g for 5 min and resuspend in deionized water. This centrifugation was repeated one more time in order to ensure a clear spore suspension free of metabolites. After the final wash, remove the supernatant and  resuspend the spores in water containing 0.05% Tween-20. Estimate the spores in this suspension by  haemocytometer or through McFarland and adjust  to the required concentration.

Good luck

Hello Georgina Pegu

The same answer to you as well.