Science topic

Microalgae - Science topic

A non-taxonomic term for unicellular microscopic algae which are found in both freshwater and marine environments. Some authors consider DIATOMS; CYANOBACTERIA; HAPTOPHYTA; and DINOFLAGELLATES as part of microalgae, even though they are not algae.
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Hello everyone,
I am going to prepare either an alginate or carrageenan hydrogels for immobilization of microalgae. The microalgae will be expected to be retrieved from the hydrogels after finishing cultivation. In that case, hydrogels need to be destabilized/dissolved. Can anyone suggest any chemicals or processes for this ? I found some information about chelating agent like sodium citrate or EDTA as chelating agent for Ca2+ ion in alginate matrix, but not sure any other agents out there are available, especially for K+ ion.
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I expect yes
📷
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Hello friends. I want to extract rubisco from spirulina microalgae does anyone have a solution? Do you have a proper protocol?
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What is rubisco
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For some reason, I had no choice but to culture my microalgae in the same LAF that was being used for fungal studies as well. Now for obvious reasons I have started getting fungal contamination in my microalgal plates. I have changed the LAF and even on subsequent re-streaking, I am unable to get rid of it. Any suggestions as to how to make the microalgae free from fungal contamination?
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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.
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I found a published photo where author writes that this is diatom species Hyalosira delicatula. But I have some doubts about such conclusion.
Unfortunatedly I do not have any photographs and the description of Hyalosira delicaula. So, can anyone familiar with the genus Hyalosira confirm or refute this conclusion?
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Roksana, thank you.
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This algae invade my culture lately (Chlamydomonas). It will caused the culture to clump and eventually die. This contaminant was motile, with more than 4 flagella observed.
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Dear Jess,
probably it is Poterioochromonas, a golden algae. You can find some information here:
Best wishes
Eleftherios
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Hello...
Two months ago, I bought pyrocystis fusiformis culture for my research. Unfortunately, no bioluminescence was observed, and the culture seemed to be deficient from the start.
Now I'm trying to buy a Pyrocystis fusiformis culture again and found a lab called PyroFarms, but before I order, I wanted to check if anyone had any past experience with them. or alternatively, if you could name any other trustworthy sources where I could buy a P. fusiformis culture, I would be grateful.
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Pyro Farms, highly recommended
Joel
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Hello everyone. Hope this question find you well.
I was recently reading about fatty acids because my master thesis is going to be about them and the most of their perspectives, like a review. And a question arose since I read in a book that there are some trans-unsaturated fatty acids found in nature (microalgae, bacteria, plant, etc), I can assume why and their reasons to be, but my question is that if these trans-unsaturated fatty acids can turn in cis fatty acids because some enviromental condition, as a reduction in the environmental stress to which it may have been subjected the sample.
I was searching for some information that can answer my question, but nothing at the moment. I'll appreciate so much your help with this. Thank you!
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Yuvraj Saxena thank you for your answer! it truly helped and brought me a little more information about it
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I found this specie in estuarine waters of Ecuador
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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.
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Good afternoon, I am working with the production of Chlorella sp. in industrial waste and I would like to know what parameters are the most optimal to evaluate in this industrial waste... If anyone has any information about it, I would appreciate your contribution.
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pH, NPK, C content, DO, moisture. If you need any more information then write back to me
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This one is a marine microalgae isolated from Bay of Bengal.
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Can you provide more clear views
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Good day All
I am busy cleaning up the microalgae strain Haematococcus pluvialis isolated from my bird bath outside. I have been battling to get rid of the Paraphysoderma sedebokerense (fungal parasite) on this strain for a very long time. Is there perhaps someone in this community with more experience who can advise me on which specific antibiotics and / or fungicides would be most effective for cleaning H. pluvialis? Your help will be much appreciated.
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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.
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This is Haochen Zhang, a MA design products student from Royal College of Art. And I’m currently working on a project about removing nutrition of flashed dairy manure. After learning some relevant materials, I realized that microalgae is a very effective means of treating wastewater, and I want to design products which can take use of microalgae (Chlorella) to help local rural residents transform the water to new resource. But now I lack of detailed and professional guidance. So I sincerely hope that I can get an opportunity to learn from you, and get more advice about this project.
Here are some questions which have been confusing me so much:
1. Will sterilization influence the effect of chlorella growing? If I want to sterilize the flushed manure water (100% sterilization), how much do I need to heat it to what temperature? And how long should it last? Can a sterile environment be achieved by simple boiling? Will high-temperature heating produce toxic substances in manure?
2. Is it necessary to re-dilution to feed chlorella with flashed manure water? Is it sufficient as a source of nutrition for chlorella? Do I need to add more nutrients? To what extent can the nitrogen and phosphorus in the water body be absorbed by chlorella?
3. What indicators should be paid attention to in the environment of cultivating chlorella? For non-professionals, is it difficult to cultivate chlorella with the water? What is the most suitable container volume for cultivating it?
4. Can the cultivated chlorella be directly used for fertilization (spraying or watering on vegetables)? Does it have a positive effect or additional economic value for agricultural production?
I sincerely hope that I can harvest precious opinions and opportunities to learn from you. Your suggestions must be very valuable to me. I hope with your help, my project can become a good design that has a practical effect on society! If possible, I hope to have more contact with you, thank you very much! !
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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?
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Despite the high quality of fish-based ingredients in aquafeeds, reported unsustainability caused by their use in aquaculture, have raised global concerns and effort for replacing them.
Among the potential candidates ( insect meal & oil vs plant meal & oil vs microalgae meal & oil : Spirulina , Chlorella, etc. ), which ones have no side effects or don’t generate new environmental impacts ?
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Microalgae considered a a noble biomass for production of high value products, food and feed, Pharmaceuticals as well as Biofuels.
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Thank you Gilbert for your valuable feedback regarding this!
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How does the F/2 medium help the growth of the Dunallila salina microalgae?
why do we use this medium?
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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.
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What is the main mechanism of wastewater treatment through microalgae in open ponds and closed cultivation? How does it work when there is no oxygen and carbon dioxide in closed cultivation?
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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.
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I'd like your advice on Pyrocystis Fusiformis growth. I purchased a culture a month ago and began cultivating it right away. Things, on the other hand, do not appear to be moving forward. There has been no bioluminescence observed yet, and the algae's growth has been poor. The culture was maintained at a pH of 8.2 and a temperature of 20 degrees Celsius with a 12-12 dark/light cycle. Cell counting under a microscope was used to monitor the algae's progress, but because I don't have much experience in the field of microbiology I wasn't sure of my count. I've attached a photo of what I see under the microscope, and I'm not sure if it's algae cells or not.
I'm hoping you could give me some suggestions on how to improve the growth of algae and its bioluminescence.
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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.
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Hey all !
I am looking for the best method to prepare microalgal slides for TEM characterization, especially as I am working with nanoparticles.
Thanks all in advance :)
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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.
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I want to know the pro and cons of this production?
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Micro algae is a better option. However, you have to cultivate the correct species. Otherwise oil yield will be low.
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Good morning, I have a question and I hope someone can help me. I have as inoculum chlorella vulgaris, with photoperiod of 12/12 hours, shaker agitation up to a volume of 500 mL and agitation provided by fish tank pump for volumes of 1 L and up.
I have observed the formation of a foam on the top of my crops, which I associate to a bacteria producing it but I have not been able to control that, any recommendations?
Also in the upper part of my containers it is forming like a green slime where my microalgae are staying and although I try to homogenize them they do not unite. Could this be associated with
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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
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Good morning, someone could help me to decipher what is causing this in my solid media, I planted chlorella vulgaris in plate count medium and in nutrient medium and I observed that some of them became fuchsia with the passing of lso days, could it be some metabolite or some microorganism?
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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
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Can I extract carotenoid pigment from microalgae (C. sorokiniana and D. salina) without freeze-drying?
I already searched and read references about drying process that we can use heat-drying method but that wasn't recommended because could cause degradation of carotenoid content. So if I don't have a freeze-dryer, can still extract the pigments from wet biomass?
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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.
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Hello,
For a project aiming to produce Spirulina production for human use, I am lokking for a pilot scale (around 10 or 20 thousand liter capacity) PBR supplier..
I will appriciate if you suggest a good company producing LED light integrated tubular PBR in Europe or Asia.
thanks alot for your help
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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.
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I recently published an article "Shining a Light on Wastewater Treatment with Microalgae" doi.org/10.1007/s13369-021-06444-3 https//rdcu.be/cE0C2 and now I would like to prepare a much shorter version of the article for publication in Frontiers for Young Minds. There are no publication charges, and this is probably the best way of communicating with the next generation of scientists. Because the article is intended for younger students the journal limits the length of manuscripts to 1500 words, 3 figures, and 5 references. The scope of the article will be determined by the team that agrees to collaborate on this venture. It is not required that authors are well versed in wastewater treatment, microalgae, or renewable energy. Rather it is only required that you have an interest in these topics and are willing to collaborate.
Can you help?
John Kilbane
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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.
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We are starting soon an in vivo trial with broilers fed diets supplemented with microalgae and insect full fat larave. We will collect blood samples and I'd like to have your suggestions about the best parameters to be analysed on blood to evaluate the oxidative and metabolic status
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Kindly check also the following useful RG link:
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I m working on biosafety evaluation of GMO microalgae, can anyone guide me which is better and acceptable method (lyophilisation or simple heat drying) for drying as i am going to use it as feed for fish? Also after harvesting pellet of algae, at what temperature i can store before drying, 4 C or -20 C, and for how long without affecting cell integrity?
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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.
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My name is Stanley please help me with the name of the microscope that I can use to view the microalgae structure and able to isolate a single strain.
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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.
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Actually for my thesis bg11 medium sample in scientific shop are unavailable. So for that any alternative to bg11 medium
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You could prepare BG11 from salts in your lab (www.uni-goettingen.de/de/186449.html)
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Photoautotrophic regime is known to be not as productive as photoheterotrophic regime, mainly due to the fact that in photoheterotrophy microalgae has both light and organic carbon availability. Is there a situation where the opposite happens? What could be the possible explanations? I appreciate your thoughts on this.
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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!
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Hello! I am trying to extract chlorophylls from microalgae, and I would love to ask how can I separate chlorophyll a from chlorophyll b? I want to get each of them separately. Thank you in advance.
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I am running a drying model and would like to have the water content of a chlorella vulgaris microalgae, before drying. Thanks. best.
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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
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Currently I and my group members was tasked on a project on using carbon dioxide as main feed for microalgae product formation. So, we decided on using Chlorella vulgaris but now we are all stuck on balancing the equation since
1. we failed to properly formulate chlorella vulgaris
2. the pathway we have chosen does not show its by products
Without those 2 we cant do economic potential analysis to see whether its profitable for production. any ideas how we should proceed? we may need to change the microbes we use thus the product and that put us back into square one. btw we are trying to get beta carotene as our main product. and the pathway is chosen from. or have we gone the wrong steps?
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You can also try "stress imposed by environmental conditions, such as , temperature, heavy metal concentration, and nitrogen availability; and also UV light stimulation.
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I am making kosaric media using artificial seawater in various salinity for microalgae cultivation. At first, I tried to filter the seawater and then add chemicals. After autoclaving, there was some deposits at the bottom of the media. I also tried to autoclave the artificial seawater and media separately, and then let both solutions cooled down and added together in laminar flow. However, once I added the kosaric media into seawater, precipitation was observed and large amount of white crystalline substance at the bottom. Is there any other way to make it?
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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.
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I want to quantify total carbohydrates in Hemerick media ( a media for microalgae growth). After I added phenol & sulfuric acid the sample becomes totally black. This problem isn't observed with artificial sea water media and the same phenol/sulfuric acid solutions.
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Hey Maximilien,
If You have a look at the components, the problem arises from nitrate. Nitrate reacts with sulphuric acid as NO3- -> HNO3 -> NO2+ + H2O.
NO2+ 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
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Hi everyone. I am measuring carbohydrate content of some microalgae. While I was making the standard curve, I noticed that the cultural media (F2 medium and ESAW medium) can also react to phenol and sulfuric acid, leading to a strong yellow color which affects the final result. Do you have any idea why this happens? And how can I solve the problem? Thanks a lot!
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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
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I know that this depends on the rheological factors of the medium, but if you know any, please let me know.
Best Regards,
Roberto Novais
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You have to calibrate according to your growth conditions.​ Make​ a​ standard​ caurve and​ compared batween OD and​ cell​ counting
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I am conducting an experiment to obtain enhanced antioxidants production in microalgae by implementing a 3x3 full factorials (three factors each at three levels) abiotic stressors. I would like to analyse my data using minitab software or jmp and obtain optimal levels of factor for maximum antioxidant production.
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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.
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I am a PhD student, working with microalgae,
i have a doubt, can microalgae utilise water soluble cellulose like CMC as carbon source?
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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)
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We are an certification body for organic farming. We also certify some producers of microalgae, such as Spirulina and Chlorella, in Asia. According to organic standards, only organic fertilizers may be used in such a system. Our certified operators claim they use fertilizers such as soybean meal, but we have more and more doubts if that is possible and plausible at all. So far, I have found a few papers confirming that nutrient uptake from organic sources is less efficient. But these papers were based on research combining mineral and organic fertilizers. So far, I have not found any publication confirming it is possible or impossible to work exclusively with organic fertilizers. Maybe the lack of such papers shows that it simply does not make sense and we are on a completely wrong track? Any reference to papers and/or answer from people who have knowledge / experience in this field are most appreciated!
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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.
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I'm trying to grow scenedesmus obliquus, chlorella vulgaris, scenedesmus sp and chlorella sp.
On Monday I inoculated 4 mL of inoculum in 4 mL of BG11 medium (prepared as 1 mL of BG11 and 999 mL of miliQ water) in separately falcon tubes. I put also agitation. The final mix was very green.
However, on Wednesday, the falcon was so clear and all the microalgae culture was dead.
I'm doing something wrong with the way of growing the microalgae?
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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.
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Image 1 and 2 (F8 P 100x). Possible diatoms. No idea about ID
Image 2. (F3 P 20x). Preliminary, order Naviculales Gyrosigma sp
Image 3 (FV 40X). Preliminary ID as Glenodinium sp
Image 4 (F9 100X) and Image 5 (imagen1). Same microalgae. I don´t have a clue about it clasification
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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.
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What is in your experience the best way to kill contaminant cyanobacteria in a green microalgae culture? Antibiotics, and in case which one do you tried?
Have you ever tried to apply variations in the physical or chemical culture conditions?
Let's me know your experience!
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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 (
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Does anyone have experience running microalgae samples in the cytometer? I would be grateful if you could share any advice on avoiding the samples getting stuck in the cytometer lines and clogging the equipment. I use an Attune NxT cytometer, and the samples are from Isocrysis galbana.
I really, really appreciate any help!
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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.
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I am working on a synthetic biology project in which I am redesigning the pathways in photosynthesis of Synechococcus elongatus to become more efficient, evaluated by increase in biomass. We want to computationally model this in R but we are struggling to find packages that can help us with this. I've looked at deSolve::aquaphy which models C and N assimilation but photosynthesis rate is an input and we want to model this also.
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I know from literature that the range is very big like between 80 - 1000 µmol/(m²*s). My aim is to find a LED light source between the range of 150 - 300 µmol/(m²*s). When I look on market ther irradiance is often given in LUX and lumen. So how can I convert his value in µmol/(m²*s). Is there an "easy" equation for it, which I can solve in Excel ?
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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.
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I need a list of laboratories please, can someone help. Thank you
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Thank you Dr. Maria for your answer. Actually, we did so, thanks a lot.
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We will use filtered tap water for large-scale microalgae production. What should be the features of the filtration device? What are the optimal water properties of tap water for the best growth of algae?
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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.
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To culture asian seabass / barramundi (Lates calcarifer) larvae, we are using rotifers as live feed, needing to be enriched with essential fatty acids (DHA and EPA):
1- Feeding rotifers with microalgae like Chlorella and enrich themby fish oil as DHA source before being fed to fish larvae.
2- Feeding rotifers with DHA and EPA rich microalgae such as Isochrysis galbana and Nannochloropsis oculata (no additional enrichment is needed).
Which one do you recommend and why?
Thank you
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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.
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Hello,
I am trying to conduct an experiment where I test the effect of different environments on the amount of oil produced from microalgae, more specifically, Chlamydomonas reinhardtii. I was planning on doing a condition wherein which I differ the amounts of nutrients the algae get. I am having trouble finding a growing medium to grow my age, I was wondering if anyone knew how I would be able to get this? Or perhaps know a recipe to make one?
Thank you
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Greetings,
we are currently working on the cultivation of the microalgae Dunaliella tertiolecta. We try to count the cells in a Thoma cell counting chamber under the microscrope. Problem is, the cells move phototactically, thus the counting of cells is inaccurate at best, caused by rapid movement of the cells under a light source.
I'm wondering if anyone knows of a method to immobilize the cells? I'd be very grateful for any suggestions!
Best regards and thanks in advance,
Marius Tölle
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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
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To know the carbon fixation rate of a microalgae culture, several bibliographic sources use formulas that consider the biomass change between two times and, knowing the carbon fraction of the biomass and with the molar mass of CO2 and C, we can obtain the carbon biofixation rate of a culture.
However, this raises a question. The variation of the algae culture occurs in the growth phase. But if I have an algae culture in a constant maximun biomass for a year (per example), the carbon fixation rate still aplies over that year?
If that volume of culture fixes X gC02 per day, could I estimate with this formula the annual fixation of C02? Or there is another way to stimate the carbon fixation per year of an algal culture of constant biomass
Thanks in advance for your time
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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.
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I'm starting to work on biofuel from microalgae but I have any idea to collect them with a simple méthod it's a work from my TFC
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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
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Question of the day!
Dear Algal Phycologists and Biotechnologists,
I have read the literature about different method of harvesting of Microalgae but I am confused which is the best method as there is no literature up-to-date calming the best harvesting method which is highly efficient. (Flocculation/Sedimentation/Filtration/ Floatation etc.)
Could you please let me know the highly efficient harvesting method up-to-date.
#research #algae #algartech #algaebiotechnology #microalgae #microalgas #cyanobacteria #diatom #algalbiofuel #phycology #phycobiotechnology #algaebiology #marinebiology #marinebiotechnolgy #biomass #energy #environment #engineering #chemistry #sustainability
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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!
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micro plastic contamination will reduce sunlight penetration and affect the photosynthesis activities and other impact. do have any report or case studies in this regard
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I´m growing Scenedesmus sp. and Chlorella sp. inoculum in a f2 culture medium opened in 2012. It's possible that this issue interfere in the growing rate of the microalgae? I noticed that these solution include vitamins, it's possible these vitamins expired?
Thanks for all the answers.
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f/2 is typically a seawater medium and Scenedesmus and Chlorella are freshwater algae - that is likely the problem.
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If it is possible, suggest me a protocol, or suggest me a researcher or lab where this kind of research is active. Particularly, the main interest is cell division rate and cell morphology change study under different climatic situations.
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Søren Busch@thank you
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Hello,
Research papers are mostly focused on lab-scale lipid extraction which most of them are not cost-effective. Also, old methods such as Folch and Bligh & Dyer are not environmentally friendly.
I wanted to know which lipid extraction method from microalgae is currently available and is being used on the pilot and industrial scale (environmentally friendly and cost-effective)?
Thank you so much,
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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.
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I have the value of Delta N for 3 producers (water column phytoplankton, sedimented attached microalgae, periphyton), primary consumers (zooplankton), tertiary consumers (different fish species).
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It is well described in the book by Brian Fry 'Stable Iostope Ecology'. Also very useful when it comes to calculation of mixing-models.
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How is citric acid beneficial to microalgae growth?
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Do you mean the citric acid need if we are using Fe in our medium?
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I tried fixing the cells, treatment with 2 % SDS but the dye is not goin inside the cells.
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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.
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Hello! I am needing some help with HPLC - I had to pick it back up after years of not doing it.
Re-learning has been going well, but I have never had to collect compound fractions and this is what I need help with.
I am performing extracts on microalgae to detect and isolate specific compounds in the UV range. My extract is generated using 100% HPLC -grade Methanol, and my eluents are 100% methanol and DDI water (all compounds have been filtered by 0.22un membrane and degassed via vacuum pump) for my flow gradient. I generated great chromatographs, spectra, etc
BUT, how do I collect specific fractions for analysis? I have multiple peaks at various times that I need to collect. I know that I can manually time it and manually collect the sample, but I need better resolution than a manual collection. I've been looking for manuals and protocols online, but the information for my systems seems to be sporadically available.
Any chemists/biologists/physicists or experienced HPLC users have some tips or resources?
System Information:
Waters 2695 Separations Module w/Column-Heating Cabinet (set to 40*C) - Hardware
Waters 2996 Photodiode Array Detector - Hardware
Waters Fraction Collector II - Hardware
C18 Column - Hardware
Empower 2on a Windows XP OS - Software
There is also a Waters 2424 ELS Detector (Hardware) , but I don't think I need this.
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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.
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I am interested in anaerobic co-digestion of algae and lignocellulosic biomass, I am uploading the related research paper, with this question.Please gi e your kind suggestions.
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Thanks for your kind suggestions.
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I need to prepare bold basal medium for the cultivation of a microalgae Chlorella species, but want to know if I can use something like e.g cobalt chloride as a cobalt nitrate substitute.
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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:
Enhanced Vitamin B12 Production using Chlorella vulgaris
This paper has been posted on RG as public full text and can bee freely downloaded as pdf file.
God luck with your research! 👍
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After searching among literature for months, I am not still able to choose a microalga strain for my PhD project. The idea is to extract as many as possible valuable compounds (such as pigments, PUFAs, polyphenols, phytosterols, vitamins) from the same microalgal biomass.
To do this, I need a strain that is being already studied and optimal conditions growth for bioproducts target are known. Also, I am not sure that selecting strains already on the market (maybe lipids or biofuels market) could be a good idea.
Which strain do you suggest?
I've found a promising strain (Stichococcus sp. KMMCC 365 but it's impossible to find to buy)
Thanks to everyone that will spend sometimes to reply me
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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.
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In a typical paper around microalgae biosensor, an increase in current was illustrated with time by chronoamperometry. But, my observations showed a decrease in current with time and I wonder what problem may occur?
Could it be cause of electrochemical method or the kind of microalgae species?
Please share me your experiment and knowledge.
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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
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I have delta N for Snail, 3 source organisms (organic matter from phytoplankton; benthic microalgae and peryphyton); primary consumers (zooplankton) and tertiary consumer is Hilsa fish of different sizes.
May I apply TP= [(deltaN primary consumers -deltaN primary producers)/3.4]+1 ( Post DM (2002) Using stable isotopes to estimate trophic position: Models, methods, and assumptions. Ecology 83:703–718)
If so, how can I calculate trophic transfer from primary producers to zooplankton, fish?
Should I use the average delN value of zooplankton as the primary consumer and the average delN value of each primary producers?
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2. You can consider a one-isotope mixing model and take into account trophic enrichment factors to estimate trophic contribution from producers to consumers.
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We're working on identifying microalgae and macroalgae strains and we plan on doing a PCR later on but we need to exract the DNA first and i can't seem to find methods that don't require CTAB, Liquid nitrogen and phenol:chloroform:isoamylalcohol. We currently do not have these components in the lab.
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hey, Salma Chahid I think we are in the same university. I can share those compounds with you. P.M if you are interested
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how to isolate the indigenous microalgae from water sample
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Hope this handbook helps.
Please check Chapter 4 - Microalgae Isolation and Basic Culturing Techniques.
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I am working on a project which requires the cultivation of C.vulgaris in a commercial fertilizer. The phosphorus form stated on the fertilizer's composition list is P2O5. I would like to know if the phosphorus can be utilized.
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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
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In my previous understanding, E. gracilis accumulates paramylon when grown in high-carbon medium, especially if grown in the dark. I have reproduced this in the past using KH medium. However, I surprisingly found yesterday that E. gracilis grown in AF6 medium with 12:12 ligh:dark cycles for almost a month showed quite high paramylon content. Can anybody point me towards literature to explain this? Is there any understanding of the dynamics of this accumulation of paramylon?
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I kind of answered my own question (partially). Will leave this here in case it is useful for someone else.
  • "Although E. gracilis was not expected to store large amounts of paramylon when grown phototrophically, we observed that phototrophic cells could contain up to 50% paramylon."
I am still interested in information about the dynamics.
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Hello, I
am trying to store microalgae at 4°C and I want to know if this has no effect on the biochemical composition
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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.
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I have exposed microalgae cells to temperature stress and measure the ROS level using DCFH, but I found that the DCFH was significantly lower in stress-treated samples compared to control. How is this possible?
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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
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Funnily, this is a very prominent topic - yet I wasn't able to come up with a good answer. So here it is: Is there a recipe for a standard nutrient-depleted medium? In many applications, microalgae are starved to promote the build-up of e.g. lipids. One solution is to just let the microalgae stay in the same (exhausted) medium and wait. But what about centrifuging the microalgae and placing them in fresh medium that is poor in nutrients? And if this is done, how should such a medium look like? Suppose we want to make sure that neigher N nor P are available, does it make sense to take a medium recipe and just omit nitrate and phosphate? We would end up with a strange medium. I would rather like to have a typical "freshwater medium with standard osmolarity and no nutrients". How does that look like? Is there a standard recipe available (that is simple to make)? I had a look at artificial lake water, which is rather cumbersome to make, it seems. Is there something simpler?
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I've been working on microalgae for some years and still I've not found documents about full scale, operating works for the production of biofuels from microalgae.
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The reason is simply because the process you mention is still far away from being economically feasible
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I am performing wastewater treatment thought different microorganisms such as microalgae and afterwards determining the lipid content in the biomass though Bligh and dyer method. Due to the COVID restriction, we are allowed to have limited time in the laboratory and therefore, a lot of samples should be stored and tested later.
I have to store my sample for cell counting and lipid determination?
For cell counting, Can I store my sample in ethanol to persevere it and count it later?
(0.3 ml sample and 0.7 ml ethanol)
For lipid: Can I store my liquid sample at -20C. and perform the lipid determination by Bligh and Dyer method later?
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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)
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We want to determine the photosynthetic rate of freshwater microalgae other than the radioactive method. kindly provide a simple protocol for measurement.
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thank you all
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I am currently cultivating C.vulgaris in Bold's Basl Medium. However, even though the medium does not meet the criteria of the Redfield ratio, the culture can still grow.
Redfield ratio C:N:P Ratio (106:16:1)
Bold's Basal Medium (N:P) (1,71 : 1 )
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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