Microalgae - Science topic

I expect yes

📷

What is rubisco

Hi,

Common growth media for microalgae generally do not support heterotrophic microbes as they lack assimilable carbon sources, and the same is true for Bold media and its 3N version. It appears that natural cell lysis with growth is enriching your algal cultures with nutrients required to support contamination. Lysis is always almost negligible, but because you are using a common culturing facility, it is making your cultures more susceptible to contamination.

As algae are very slow-growing, even minute contamination is supposed to outgrow soon. Thus it is important to analyze your system and practices in more depth to suggest some measures and tricks to avoid any contamination.

All the best for now.

Roksana, thank you.

Dear Jess,

probably it is Poterioochromonas, a golden algae. You can find some information here:

and here:

Best wishes

Eleftherios

Pyro Farms, highly recommended

Joel

Yuvraj Saxena thank you for your answer! it truly helped and brought me a little more information about it

Of course not! Strictly speaking, it is a nomenclatural and spelling error. But in the case of Scenedesmus, Chlorella, Chlamydomonas, Oedogonium, and some genera that begin with compound Latin letters (Oe, Sc, Sch, etc.), this spelling was used to avoid confusion with other genera in the floristic lists. Sorry, I should have written S. indicus and S. producto-capitatus.

pH, NPK, C content, DO, moisture. If you need any more information then write back to me

Hi Thais, thank you for answering. I have streaked too many plates to count. I look for a colony under the microscope, transfer it but a few days after, I have contamination again. I suspect I am transferring the fungi spores too. H. pluvialis is quite a slow grower. I have good colonies after a week but I suspect that they probably get contaminated at day 2 - 4. The fungi grows and spreads quicker than I can get colonies. I know I can buy a clean strain but I really want to get this right and clean my own. I can see it is a good strain with large cells and very well acclimatized already.

Arne Andersen Thanks for your suggestion, it seems to be a more efficient and practical way. One thing I'm not sure about though is whether it takes longer to degrade cow dung water by the growth of saprophytes than when using microalgae?

Check https://www.fao.org/3/y3781e/y3781e.pdf

Thank you Gilbert for your valuable feedback regarding this!

This medium contains abundant levels of N and P (also Si for diatoms and other Si-requiring species) along with a comprehensive set of trace metals and vitamins. The N:P:Si ratio is close to the classical Redfield ratio. As such, the medium is very effective in supporting the growth of most algae. Another similar medium is L1, which has similar recipes with small modifications of trace metal composition.

I agree with Vit, in open ponds, there is CO2 whereas in closed at the beginning they will use organic carbon, but if there is light they produce oxygen and co2 in the dark phase o photosynthesis, so you will have mixed metabolism. Microalgae are supposed to remove nitrogen and phosphorous from wastewater, but they can also remove carbon depending on growth conditions.

Barry C Smith F/2 media was utilized, along with a quite bright LED light (5000 lux measured using a lux meter app on the inoculum surface). The culture volume is 4L, while the inoculum volume is around 6L.

Dear Maysan Nashashibi , thanks for sharing this interesting question. It is always a good idea to use the searching box into RG website, often you will get some very valuable informations such as those provided by Aarif Shah . Specially in the first link you can see a good TEM sample preparation protocol for bio-samples with nanoparticles.

That is a generic protocol, and it must be adapted to each sample and to what you want to characterize. For example, it will be different if these nanoparticles (NPs) are in the liquid media, on the microalgae´s surface, or dispersed into the microalgae´s tissues. Problems associated to NPs aggregation will probably happen in the two first scenarios, when the NPs are dispersed into the liquid media and by evaporation of it, they end up on the surface of the algae. Some NP´s could be attached to the algae surface and evaporation of the media could add more from the media. If you are interested about the NP´s already bonded to the microalgae surface, and there are NP´s dispersed into the media around, it would be useful to remove the algae from the media, wash them up with some clean media (bear in mind that this clean media should not remove the bonded NPs from your algae), then dry the algae as explained in the previous links or if it is available, use a TEM cell for liquid samples. Of course, the microalgae must be thin enough to work in transmission mode as pointed by Mohammed Amer Shaheed .

If the microalgae have the NPs inside their tissues, the chances of aggregation by drying are very few, the algae tissues will keep them apart, so that if you finally observe aggregation of NPs, it means -generally- that they were already aggregated before the drying process. Once more, Mohammed advice is key, you will need a very thin section of your algae to let the electrons go through the sample and forming a image. You can get such thin sections embedding the sample into a resin and cutting it with an ultramicrotome. Otherwise, you could use a FIB (Focused Ion Beam), a very focused and thin beam of ions to cut a lamela (a very thin section) of your sample, which is then placed on a TEM holder grid with the help of micromanipulators.

TEM works by transmission, transmission of electrons through the sample, so depending on the density of each part of your sample (the value of Z, the atomic number of the atoms on each part) you will see more or less contrast. For example, if your NPs are metallic, they will be denser than the organic tissue of your microalgae, and a good contrast will be expected. But if your NP´s are light, say carbon NP´s or Si NPs the contrast with the organic matter will be smaller.

It could be very useful, before to go to the TEM, to observe the samples with an optical microscope (may be a confocal, looking for some fluorescence from the NP´s), or a faster option, to use a SEM, specially if you are studying NP´s on the algae surface. Normally you would need to dry up your sample as with TEM, but here -with SEM- you don´t need to worry about sample´s thickness too much. SEM often requires conductive samples or coating your sample with a very thin conductive layer (gold, carbon, iridium...), but modern equipment also let us to use the sample uncoated, thanks to charge removal methods.

Hope this helps. Good luck with your research work and my best wishes.

Micro algae is a better option. However, you have to cultivate the correct species. Otherwise oil yield will be low.

It may due to the saponins produced by Chlorella. Generally they exhibits high phytochemical content especially n log phase. please subculture the same and if the property reappear go for saponin test

Hello Alejandra Vásquez

Maybe your plates are contaminated because changes in pH or growth of the microorganism may lead to a change in colour. Also, may I ask what agar are you using and exactly how long have these plates been inoculated. If your cultures were green at first and then turned pink, then it just means that the culture had become old.

To be sure, I suggest you cut a piece of the agar and grind it in some distilled water. Then observe the contents through a microscope. You might get an idea of what happening in case its a contaminant.

Best wishes,

Ritesh

As many people have answered, yes you can.

If you want to see the amount of carotenoids per weight, you can determine the dry weight per culture medium, and separately determine the amount of carotenoids in the wet biomass per culture medium.

Thank you for all of yours suggestions, Schott proveide really high tech products but it is really expensive. BBI is no longer produce reactors in pilot or industrial scale. I didnt know algoliner, I will contact them. Thnaks again.

Jacob de Feijter, thanks for your interest in collaborating on this topic. The task has already begun and correspondence among collaborators is taking place via email. You may contact me at kilbane@iit.edu and we welcome your involvement.

Kindly check also the following useful RG link:

I recommend freeze-drying. It preserves all the vitamins and other heat-sensitive micro and macro nutrients of the algae if you use them for fish as feed. If you use freeze-drying, you can store algae in -20 just after harvesting.

I would recommend an inverted microscope with sedimentation chambers, so you won’t have to squeeze your algae between slide and cover glass. This also means you can use magnifications of 630 times, maybe even 1000 with immersion oil. Use both live samples and lugol preserved ones if your specimens are too mobile.

You could prepare BG11 from salts in your lab (www.uni-goettingen.de/de/186449.html)

Thank you, Maryam Fath ! Do you have some references on this? Could you please share the link to the papers with me? I appreciate it!

Also see kindly the following link: https://graduateway.com/separation-of-chlorophyll-a-chlorophyll-b-and-beta-carotene-by-paper-chromatography/

Hello. From my experience in the lab, centrifuged microalgae sludge from a 3L Erlenmeyer flask could accumulate a fresh weight of about 2-3 grams and when fully dried would give a weight of about 0.6 - 1.0~ grams. This is a rough estimate as values can vary based on treatment conditions as well. Hope this helps with your project

You can also try "stress imposed by environmental conditions, such as , temperature, heavy metal concentration, and nitrogen availability; and also UV light stimulation.

hello dear,

i think you can first filter your seawater, then add your desired chemicals might be these chemicals react with each other i am not sure but i heard from my friend, he face such this type of problem.

Hey Maximilien,

If You have a look at the components, the problem arises from nitrate. Nitrate reacts with sulphuric acid as NO₃^- -> HNO₃ -> NO_2⁺ + H₂O.

NO_2⁺ reacts with phenol to nitrophenol, which absorbs in the UV-region and looks dark-greenish.

I was facing the same problem and I tried to solve it as follows (please consider citing me as I cannot find any comparable method):

I measured aqueous nitrate standards and found that they have a plateau around 490 to 510 nm, which is bad as it increases the absorbance exactly at the point, You want to measure the carbohydrates. When nitrate is consumed during the culture, there is a time-dependent absorbance decrease, which renders the results useless.

But there is good news: a plateau means that the slope (1st derivative) in this region is 0, so if you take the smoothed 1st derivative of the spectrum and read the value at 503 nm instead of 490 nm (steepest slope for glucose standards for me), You will get the carbohydrate value You want without the influence of nitrate (since its slope adds 0 to the steepest slope of glucose).

Of course, this implies that You do not use the calibration A(c) = a * c(Gluc) + b as for standard Lambert-Beer-Bouguer, but you need to calibrate dA/d\lambda = a * c(Gluc) + b, which is in perfect accordance with the Lambert-Beer-Bouguer law.

I verified this method in our lab: Nitrate has no influence on the kinetics of the reaction and when I calibrate with aqueous (non medium) glucose standards and measure a glucose standard in Zarrouk medium (similar to Yours), I get a recovery rate of > 95% to 100%.

So, I can assume, this Derivative-Spectroscopy-Method is adequate, but you should post a picture of your spectra (nitrate/glucose in water & glucose in medium, full spectral range) because the assay is very sensitive when it comes to reagent ratios. With this method I fixed the nitrate problem for us, but there is another one. In supernatants from real algae cultivations I get peaks between 500 and 600 nm and since they are not a plateau, they lift the slope of glucose at 503 nm, which dramatically affects my results (lowers them).

I tried to overcome this by taking the smoothed second order derivative at 490 nm (second order derivative is minimum at the top of peak), but since I do not know what causes the colour between 500 and 600 nm I cannot exclude the influence of the colour by another hidden peak below the glucose peak. The influence of nitrate is also 0 for the 2nd derivative since the second order derivative of a plateau is fortunately 0 as well. To sum it up, the new calibration at 490 nm would become d²A / (d\lambda)² = a * c(Gluc) + b (also Lambert-Beer-Bouguer) - but I cannot be certain that this is working in real samples which cause an absorbance at 500 to 600 nm after the reaction with phenol and sulphuric acid (only with both, not with one of them alone).

I've not found anything mentioning the influence of nitrate in the assay except for this (http://www.bioline.org.br/pdf?ja06013), which is soley about nitrate.

I'm really in a need for help what causes the absorbance between 500 and 600 nm, since I've tried everything (various salts, protein (BSA standard), oxidation with persulphate, acidic pre-hydrolysis, cooking, performing the assay with algae biomass still in the sample), but I was completely unable to mimic the additional peaks. I'm growing desparate and depressive about this because I cannot find any literature regarding this. The assay was just not intended to be used in saline solutions and on top of that, nitrate is impossible to remove except for dialysis (cost, effort, etc...).

Maybe we (and I hope experts) can collaborate on this problem. If someone wants to contact me, just google my name + "OTH AW". I have Excel- and Python-software for smoothing, derivation if somebody is not into programming.

Hope this helps, at least for the moment.

Best regards

Niklas Zell

Hello Ruqian,

Saeed and Anthony are correct. It is best to harvest the cells first, wash the pellet and re-suspend in for e.g distilled water. However, I believe it would be a better approach to dry your sample first and then use the dry biomass for the test. It should give more accurate results.

Kind regards,

Ritesh

You have to calibrate according to your growth conditions.​ Make​ a​ standard​ caurve and​ compared batween OD and​ cell​ counting

You would first need to do a screening study with your 3 factors in jmp, I would suggest a 2X3 (8 runs) with centre points to account for curvature rather than a 3X3 (27 runs). Use the screening study to remove factors that are not significant. Check for the validity of your model by use of graphical methods (residuals and a normal plot) and numerical methods (ANOVA and lack of fit). Once the model is shown to be sound, fit in a linear model based on your significant factors - this linear model can be used to do a method of steepest ascent if your initial screening didn't identify the region of maximum yield - After this, augment your initial screening study with axial points and carry out a response surface study using a central composite design (CCD) or a Box-Behnken design (depends on your factor ranges). The response surface method (RSM) will give you your optimum values by analyzing the prediction profilers, setting desirability to maximize yield and reading response surface curves or contour profilers. Finally, fit in the second-order model.

Dear Anand,

Yes there are different groups of microalgae that are capable to ferment cellulosic biomass for their growth, such as Aurantiochytrium sp. or other "fungal-like" protists like Pythium irregulare (capable to ferment plant roots).

I don't know in detail for CMC but there are many microalgae that can do this (search for extremophile algae too)

It is an interesting question. We are not sure to get full yield with organic inputs. But along with biostimulants, organic inputs could help in realising good yield potential.

Dear Noelia

I disagree with Chandan because i don't think that your algae are dying because of nutrient depletion, because in your medium you don't have any.

It's a matter of fact that you are cultivation in milliQ water, because you are diluting your BG11 1000 times, so all the nutrients are almost zero.

For green algae you should inoculate in not diluted BG11, in that way you will not encour in any nutrien starvation.

If you want a "lighter" medium i suggest a BBM medium.

Best regards.

The first photo is likely a Paulinella, a type of thecate amoeba, not an algal species. The second is likely a Pleurosigma or Gyrosigma species. The third photo is likely a Scrippsiella species. Photos 4 and 5 depict cells of some kind that did not preserve well with your fixative or if live were damaged by the heat from the microscope's light.

Hi Laura

Have you thought using hydrogen peroxide? cyanobacteria are usually more sensitive to hydrogen peroxide than eukaryotic microbes and H2O2 is often used to prevent cyanobacterial blooms in enclosed water bodies (

Hi Karla. Do you use nuclear isolation buffer? What helped me was that I observed it under the fluorescence microscope before taking it to the cytometer. So I adjusted the buffer according to the quality of my nucleus, analyzing if they were aggregated or integrated, for example.

Try BiGGR

dynamicFBA

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sybilDynFBA

Have fun and Happy Research!

Dear Halil,

I can figure out two different issues in your question. First is the light source which can provide you the required range of PFD for your experimental work - i.e. 80 - 1000 µmol/(m²*s), and the second is how one can measure/convert the common intensity units.

Commercially available LED sources generally don't provide that much PFD (80-1000). I would suggest you to look for very high wattage LED sources or customise your own light system, as I did in my studies ( ) to get the desired intensity.

Thank you Dr. Maria for your answer. Actually, we did so, thanks a lot.

you can disinfect the water with bleaching powder that is a very common chemical for treating swimming pools. 1 g/L and after that aeration for 24h. also, you can use O3, UV, or filtration techniques.

As there was no answer to this question, we tested both methods. We noticed that rotifer enrichment through feeding them with algal diet is more suitable. Not only less time, energy, and budget were needed, but also less contamination in rotifer tanks occurred.

@bethany hunt

Hello Marius,

When i need to count motile green algae such as Dunaliella or Tetraselmis i use a diluted solution of iodine tincture, just few drops in your sample, but don't exceed or you will have a too bright color.

Otherwise you can prepare the classic lugol solution or a 5% formaldehyde solution

Better to estimate the biomass production of algae per unit time and per unit area, the calculate the carbon content of the biomass. But still, estimation of the amount of carbon that could be decomposed and released into the atmosphere must be calculated. In short, the humification and decomposition coefficients of algae must be considered. The plant that fixes quickly and released carbon by simple decomposition could not be considered a potential carbon sequester.

Bonjour Premice, je me permets de te répondre en francais pour plus de simplicité. Pour la récolte de microalgues à échelle de laboratoire (autour de 5L), il est préférrable de faire usage d'une centrifuge. Mais si c'est plus que cela, il vaut mieux utiliser plusieurs techniques en séquence: sédimentation/floculation ensuite centrifugation.

A noter que si tu cultives des microalgue qui sont moins de 5um, il serait mieux d'utiliser la floculation en modifiant le pH ou d'utiliser des produits chimiques pour agglomerer les cellules. Une fois les cellules ont sedimenté, tu peut utiliser la centrifuge.

Salutations,

Ritesh

Dear Mohammad Sibtain Kadri , harvesting microalgae is not a straightforward process as such because it varies on numerous factors. One of the most important aspect to consider in this process is the dewatering of the microalgae slurries while limiting damage to the cells, biomass loss and limiting costs. All of the techniques you listed above are useful depending on the type of culture, volume of biomass production, the organism being cultured, desired end product, among many others but a crucial one would be costs involved (how far can investments be made and how much can be spent on energy driven processes).

A simple laboratory experiment might lead to the use of centrifugation directly as indicated by Aradhana Srisivasailam or a combination of them (sedimentation prior to centrifugation). However, as you scale up the production, the techniques then tend to be expensive due to energy consumption.

For my experiments, I make use of auto-flocculation followed by centrifugation at 5000 rpm for <10 min with microalgae Rhexinema paucicellulare, which tends to agglomerate and thus become heavy and sink. Smaller microalgae such as Chlorella sp. will need more intense centrifigation and flocculation might require to be chemically induced.

Therefore, I suggest you decide the most appropriate harvesting process for your experiment by conducting an elimination process based on the various aspects.

I suggest you look into the following papers:

1) Conversion of microalgae to biofuel

doi:10.1016/j.rser.2012.03.047

2) Microalgae: the next best alternative to fossil fuels after biomass. A review

doi:10.4081/mr.2019.7936

3) Harvesting microalgae by flocculation–sedimentation

Hope it helps!

Please take a look at the following useful RG links.

f/2 is typically a seawater medium and Scenedesmus and Chlorella are freshwater algae - that is likely the problem.

Søren Busch@thank you

Currently, the solvent extraction technique is most widely used for lipid extraction as they provide the highest lipid recovery. Adopting the mechanical methods, although environmentally friendly and economic, is not considerable due to poor recovery and the possible dilapidation of lipids. Solvent-free methods appear significant only at the lab. scale however further investigations are required for minimal use of solvents for large-scale commercialization.

It is well described in the book by Brian Fry 'Stable Iostope Ecology'. Also very useful when it comes to calculation of mixing-models.

Do you mean the citric acid need if we are using Fe in our medium?

I had no problem with Euglena and Chlorella cells when fixed or dead, by doing ethanol fixation and incubating with DAPI. For Chlorella vulgaris, for instance, if you want to stain both nucleic acid (blue) and polyphosphate (yellow), I used 100 µg/mL DAPI 10 min incubation following Ethanol 1:1 fixation. For Euglena gracilis, 2 µM of DAPI gave good results after a similar incubation. If you want to stain live cells, you will need permeabilization, I have been using it in either fixed/permeabilized cells (with ethanol) or as a membrane integrity dead staining.

One thing you will need to do is verify that the HPLC and fraction collector are "aligned". Some HPLC systems specify tubing lengths from the detector(s) and fraction collector, suggesting their software compensates for the delay time. This link (https://www.waters.com/webassets/cms/support/docs/71500012102re.pdf ) page 86, makes reference to a delaying time (I realize this is for a different fraction collector than yours, but the concept still holds).

I think you can choose the peaks for collection from a prior run.

Thanks for your kind suggestions.

Dear Dewald De Villiers many thanks for your very interesting technical question. Yes, cobalt(II) nitrate can be replaced by cobalt(II) chloride in Bold’s Basal Medium. This has been reported in the following research article:

This paper has been posted on RG as public full text and can bee freely downloaded as pdf file.

God luck with your research! 👍

For doctoral research, I don't think it would be a good idea to select any strain which has already been explored. I would suggest exploring new strains from the natural pools of that specific area where you are pursuing your doctoral research.

Dear Dr. Berthuel

Thank you for your reply. In fact, I used the stirrer in my test but the the trend of signals did not change. I use an autolab for receiving chronoamperometry scan, the working electrode is glassy carbon with Ag/AgCl ref.electrode and I use BSA-Glutaraldehyde matrix for immobilizing microalgae cells. Here is a paper with similar work, it indicates an increasing slope in chronoamperometry and I gained exactly reverse.

I would be grateful if you could help me for solving this problem.

This is the mentioned article: Chlorella sp. based biosensor for selective determination of mercury in presence of silver ions

doi: 10.1016/j.snb.2012.02.009

2. You can consider a one-isotope mixing model and take into account trophic enrichment factors to estimate trophic contribution from producers to consumers.

hey, Salma Chahid I think we are in the same university. I can share those compounds with you. P.M if you are interested

Hope this handbook helps.

Please check Chapter 4 - Microalgae Isolation and Basic Culturing Techniques.

The answer is very simple. No, they cannot, because fertilisers do not contain P2O5.

To read more about this, check this Editorial: Lambers H, Barrow NJ (2020) P2O5, K2O, CaO, MgO, and basic cations: pervasive use of references to molecules that do not exist in soil. Plant Soil 452: 1-4

I kind of answered my own question (partially). Will leave this here in case it is useful for someone else.

From:

I am still interested in information about the dynamics.

I tried this method, and I was successful. I have kept a mass of semi-dried (4-6 hours within silica desiccator) Chaetoceros sp. at 4 degree C in the refrigerator, and I was able to reestablish the live culture of the species from those semi-dried refrigerated pellet. Please note that, the pellet must not be full dried. In that case, re-culturing is impossible.

Dear Bahram Barati,

There are chances of decreased expression of ROS in the system under the stress condition if this is the result of the repeated standardized experiment.

But, decreased expression of ROS also tagged with other parameters such as involvement of various antioxidant enzymes, the incubation time factor, methodology of the experiment, etc.,

you can run parallel experiments regarding the antioxidant enzyme studies and GSH in order to confirm with the obtained result (decreased ROS in stress condition than Control).

All the best

Also check https://www.sciencedirect.com/topics/engineering/nutrient-limitation

The reason is simply because the process you mention is still far away from being economically feasible

For cell counting you can just add 1-2 drops of lugol solution to a 10 ml sample and store it.

For lipid determination, I recommend you to store your Microalgae pellet in the own extraction solvent with an antioxidant (BHT is commonly used) and inert (N gas) atmosphere in the freezer (-20-40ºC is OK)

thank you all

Hi, you can find different understanding in literature of what is meant under Redfield ratio. Correctly it is the generally observed ratio of 106:16:1 C:N:P in oceans (A. Redfield 1934) as you mention it. But sometimes people refer to "Redfield ratio" as any C:N:P, which is confusing, of course. Many organisms (and your Chlorella