Cyanobacteria - Science topic

The concept of vernalization as it's traditionally understood in higher plants "A cold-induced transition from vegetative to reproductive growth", doesn't fully apply to algae, including cyanobacteria and macroalgae, because their life cycles and reproductive processes differ significantly from those of higher plants and they don't experience true vernalization as defined for higher plants. However, algae can still exhibit cold-induced responses that affect their growth and reproduction, which might resemble vernalization in some ways but are not the same.

But in some brown and red algae species, cold temperatures are crucial for the transition between life cycle phases (e.g., from the diploid sporophyte to haploid gametophyte stages). The cold acts as an environmental signal that synchronizes reproduction.

May consider using Permai fluorescence dye.

salam, Thank you very much, Dear Qasim I added your detailed reply in the question part, to other researcher's usage. I think to use a finer mesh size, had better because of the diatoms. Thank you for your kind reply. best wishes Jafar

This seems a common issue. Also, in the paper you mention (Kearney et al., 2022) it was stated that cyanobacteria need helper bacteria so bacterial growth on this medium is somewhat desirable. As for the organic carbon content - natural seawater does contain some dissolved organic carbon, especially if it was procured from a coastal region. This can be reduced by "aging" the water (https://www.researchgate.net/post/Why-must-seawater-be-aged-for-use-in-media-prep). Using a high purity agar (Noble agar, electrophoresis agar) can also help minimize the carbon content.

Hello Ela,

I am not sure about the statistical tests, but the answer given above sounds sensible. I do have some thoughts on why no blue-green algae were found yet. Most systems are dominated in the spring by green algae, diatoms, and flagellates. Blue-green algae typically tend to proliferate as temperature increases and phosphorus concentrations decrease because they can compete better for nutrients when phosphorus is limiting. We usually see blue-green algae in mid-July through September when looking at seasonal dynamics in lakes, unless the lake is very eutrophic (then there are some type of blue-green algae year round). The lakes where we study blue-green algae blooms (HABs), we find that the peak biomass occurs in August. Other things that may contribute to what you might be seeing early in the season have to do with storms, wind and wave action, and if there are secondary sources nutrients entering your system via land use. Let me know if you would like to discuss this further.

Hello Alexis, thanks for the new image. Your image was captured using a backscattered electron detector with a 15kV beam. Your biological material is probably the wispy "gauze" like substance on top of the "particle layer". You may be able to see this more clearly by using a secondary electron detector at 2-5 kV. I see you used a Phenom - it may not have an SE detector. You could try dropping your voltage with your BSE detector but image quality may drop off. The "particles" are 300-500 nm so too small to be microalgae (Chlorella is 5 microns). They could be bacteria or an inorganic substance. They are quite easy to see - have you fixed and gold coated this sample. EDX should help. The sodium peak should have a larger chloride peak if there is a lot of salt. It can crystalize as cubes or show a dendritic pattern.

Making a guess here since I haven't worked with bacteria since late 1980s--the genus name Microcystis suggests a bacterium with an unusually tough cyst wall (I vaguely remember cyanobacteria in general as being very tough micro-organisms). Its good you've tried additional proteolytic enzymes; you may also want to try enzymes that are especially efficaceous against petidoglycans; perhaps sugar polymer constituents are interfering with the purification process. I would guess a web search for your problem would turn up something more reliably useful!

It seems very concentrated. Try adding it in the range of 100-200uL/L.

Extracting DNA from Cyanobacteria, which are gram-positive bacteria, can be a bit challenging, but it's possible to do it on-site with the right equipment and protocols. Here's a general outline of the steps involved in extracting DNA from Cyanobacteria:

Cell harvesting: Cyanobacteria can be harvested from various sources such as soil, water, or biological samples. For on-site extraction, you can use a sterile loop or swab to collect the cells from the surface of a culture or sample.

Cell lysis: The next step is to break open the cell membrane and release the DNA. You can use a lysis buffer, such as Tris-EDTA, to break down the cell membrane and release the DNA. You can also add a detergent such as SDS to help break down the membrane.

DNA release: After lysis, you can add a DNA release agent such as proteinase K to break down any proteins that may be binding to the DNA. This will help to release the DNA from the cellular debris.

DNA purification: The DNA released from the cells will be in a mixture of cellular debris and other contaminants. You can use a purification method such as phenol-chloroform extraction or ethanol precipitation to separate the DNA from the contaminants.

DNA concentration: After purification, you can use a DNA concentration buffer, such as ethanol or isopropanol, to precipitate the DNA and concentrate it.

DNA analysis: Finally, you can analyze the DNA using various techniques such as PCR, restriction digestion, or sequencing.

Here's a simple protocol for on-site DNA extraction from Cyanobacteria:

Collect the Cyanobacteria samples using a sterile loop or swab.

Add 500 μL of lysis buffer (Tris-EDTA, pH 8.0) and 100 μL of 20% SDS to the samples.

Vortex the mixture for 10-15 minutes to break down the cell membrane.

Add 50 μL of proteinase K (10 mg/mL) to the mixture and incubate it for 30 minutes at 50°C.

Add 500 μL of phenol-chloroform to the mixture and vortex it for 10-15 minutes.

Centrifuge the mixture at 10,000 x g for 10 minutes to separate the DNA from the contaminants.

Collect the DNA pellet and wash it with 70% ethanol.

Centrifuge the DNA pellet again at 10,000 x g for 10 minutes to remove the ethanol.

Dry the DNA pellet for a few minutes and then re-suspend it in 100 μL of TE buffer (Tris-EDTA, pH 8.0).

Analyze the DNA using PCR, restriction digestion, or sequencing.

RiPPs, or ribosomally synthesized peptides, are a type of natural product that are synthesized by cyanobacteria, which are bacteria that can photosynthesize. These peptides have a wide range of physiological functions, including:

Antibacterial activity: RiPPs can inhibit the growth of other bacteria, making them useful for protecting cyanobacteria from competition and predation.

Antifungal activity: RiPPs can also inhibit the growth of fungi, which can help cyanobacteria compete for resources and space.

Antiviral activity: Some RiPPs have been shown to have antiviral activity, which can help protect cyanobacteria from viral infections.

Antioxidant activity: RiPPs can act as antioxidants, which can help protect cyanobacteria from damage caused by reactive oxygen species (ROS).

Signal transduction: RiPPs can act as signaling molecules, influencing various cellular processes such as gene expression, metabolism, and cell growth.

Defense against environmental stress: RiPPs can help cyanobacteria adapt to environmental stressors such as high light, temperature, and drought.

Role in photosynthesis: Some RiPPs have been shown to play a role in the regulation of photosynthesis, such as inhibiting the activity of photosynthetic enzymes or regulating the expression of photosynthetic genes.

Overall, the physiological function of RiPPs in cyanobacteria is to provide protection, regulation, and adaptation to the cell, allowing it to survive and thrive in its environment.

I think that the best efficiency will be use of gamma rays

For Cyanobacteria I would recommend taking a course at The University of South Bohemia at Cesky Budejovice, department of Botany. For freshwater in general you could try https://www.ceh.ac.uk/training/freshwater-phytoplankton-identification. Good luck!

Ah, my fellow researcher Sydney Brown, a question of considerable importance indeed! Culturing cyanobacteria is a delicate art, and maintaining nutrient conditions over an extended period requires finesse. Now, allow me to share some insights.

In the realm of sterile cyanobacteria cultures, the challenge lies in sustaining nutrient levels as these fascinating organisms flourish. I would propose a meticulous approach to ensure both longevity and sterility.

Firstly, consider implementing a continuous nutrient replenishment system. Picture a well-designed apparatus that gradually introduces fresh nutrients into the medium as the cyanobacteria consume them. This way, you Sydney Brown maintain a harmonious balance without disrupting the delicate equilibrium.

Furthermore, explore the potential of encapsulated nutrient reservoirs. These could release essential compounds gradually, providing a sustained supply over an extended period. Think of it as a sophisticated cyanobacterial buffet, ensuring they dine at their leisure without exhausting the nutritional feast.

Now, let's delve into the realm of maintaining sterile conditions. Employ advanced filtration techniques, perhaps incorporating cutting-edge membrane filtration systems. Such measures would safeguard against contaminants, preserving the sanctity of your cyanobacterial haven.

Consider the implementation of controlled environments with rigorous monitoring. Picture a realm where sensors and automation dance in unison, ensuring that the conditions remain pristine. Embrace technology as a guardian, watching over your cyanobacterial community with unyielding vigilance.

Some interesting readings are:

In conclusion, my esteemed colleague Sydney Brown, the key lies in blending innovation and precision. Craft a symphony of nutrient replenishment and sterile guardianship, and your cyanobacteria shall thrive in a realm untouched by the ordinary constraints. May your experiments be as boundless!

Dear Jakub Hryc In the similar question asked here https://www.reddit.com/r/botany/comments/17p1ehs/change_lipid_component_from_galactolipids_to/ there is a good reference to https://academic.oup.com/pcp/article/59/6/1128/4990989?login=false where the message is not so much that phosphate is limited (which is sheer nonsense because all (cell)membranes of animals, included us humans, and most of the bacteria are constituted of phospholipids) but “these two lipids [the galactolipids MGDG and DGDG] are important for maintaining chloroplast morphology and for plant survival under abiotic stresses such as phosphate starvation and freezing”

Best regards.

You might be able to contact Professor Komárek and request the paper at his webpage: https://www.phycology.cz/people/ji%C5%99%C3%AD-kom%C3%A1rek

or go to https://www.phycology.cz/contact and then press 'People' and you will find his page.

Uniprot is a good website for researching specific proteins. For example, you can type phycoerythrin and it will pull up all proteins with that name. Each organism will have a separate entry. Then you can look at data that has been generated by other researchers on that specific protein, including size, function, sequence, etc.

Hi, please check this recent paper regarding flexible CO2 patch sensor:

-N BG 11,Because is is osscillatoriales,because -N will allow the heterocyst to appear much clearly

Dear Nilamjyoti,

in one of our studies we used the sulfamate-biphenyl method to isolate/detach and quantify alginate from Pseudomonas aeruginosa which might also work for cyanobacteria:

Supplementary information: https://doi.org/10.1371/journal.pone.0088368.s006

Good luck

Michael

See "Another researcher has claimed my publications" in https://explore.researchgate.net/display/support/Authorship for instructions how to correct this. But unlike this help page suggests, most probably it was not the other author himself who wrongly claimed authorship, but ResearchGate's automatic algorithm wrongly identified him as the author and assigned your publication to his profile. This is a frequent problem in ResearchGate in case of similar names.

The choice of method for the isolation and mass culture of freshwater algae and cyanobacteria depends on various factors, including the specific species of interest, available resources, and the scale of production required. Here are some common methods used for isolation and mass culture:

Isolation of Freshwater Algae and Cyanobacteria:

Mass Culture of Freshwater Algae and Cyanobacteria:

It's important to note that different species of algae and cyanobacteria may have specific requirements for growth, and the choice of culture method should be tailored to suit their individual needs. Additionally, water quality, nutrient availability, and potential risks of harmful algal blooms should be considered when selecting a mass culture method. Regular monitoring and quality control are crucial to ensuring successful and sustainable mass culture of freshwater algae and cyanobacteria.

Nematodes can be a COMMON contaminant in cyanobacterial cultures and can negatively impact the growth and health of the strains. To get rid of nematode contamination in cyanobacterial cultures, you can follow these steps:

Remember that preventing contamination is always better than trying to eliminate it. Maintaining aseptic techniques and regularly inspecting your cyanobacterial cultures can help minimize the risk of nematode contamination and maintain healthy cultures.

Algae Type: Both green and blue-green algae can potentially be used to create an extract for germinating rice seeds. Both types of algae contain beneficial nutrients and growth-promoting substances that can support seed germination and early seedling growth. The choice between green algae and blue-green algae may depend on the availability of the algae culture or extract and any specific properties you may want to explore. Green algae are typically easier to find and culture, while blue-green algae (cyanobacteria) are known for fixing atmospheric nitrogen, which can benefit plant growth.

It's worth noting that the success of using algae extract for seed germination may vary depending on factors such as the quality of the algae extract, seed variety, and environmental conditions. Experimentation and observation will help you determine the effectiveness of algae extract for germinating rice seeds in your specific situation.

I sugget you consult the book, " Standard Methods for the Examination of Water and Wasrewater". The latest edition is available on line as of January 2023. The book is jointly published by the "American Public Healthe Association" and the American Water Works Association".

Hello Vladimir,

1. Chroococcus

2. Microcystis

3. Chroococcus?

4. Microcystis

5. Chroococcus?

6. Merismopedia

7. Chroococcus?

I'm not totally sure with images, 3, 5 & 7, but Chrrococcus would be what I would go with.

I hope this helps.

Cheers,

Adam

An Efficient Protocol for Long-Term Preservation of Cyanobacteria Mayashree B Syiem* and Amrita Bhattacharjee Department of Biochemistry, North-Eastern Hill University, Shillong 793022, Meghalaya, India.

I could find this research article on long term preservation of cyanobacteria. Please refer

Ideally, you should use FACS, or if you must, microscopy counting.

You can do calibration curve of the above against OD750nm and use this calibration

There is some evidence to suggest that the growth of cyanobacteria undergoes complete cessation under conditions of extremely high light intensity, depending on the species and strain of cyanobacteria, the duration and frequency of exposure, and the availability of nutrients.

Cyanobacteria are photosynthetic microorganisms that can adapt to various light conditions by adjusting their photosynthetic apparatus and metabolism. However, when the light intensity exceeds their capacity for light harvesting and utilization, they can experience photoinhibition, oxidative stress, and damage to their photosystems and other cellular components (1),(2).

Some studies have reported that cyanobacterial growth can be completely halted at very high light intensities, such as those exceeding 1000 μmol(photons) m-2s-1. For example, a study by Muhetaer et al. (2020) found that two cyanobacteria species, Pseudanabaena galeata and Microcystis aeruginosa, showed a significant decline in growth rate and biomass production when exposed to 600 μmol(photons) m-2s-1 for two or eight days (3). Another study by Yoshikawa et al. (2021) found that the cyanobacterium Synechocystis sp. PCC 6803 showed a complete cessation of growth when exposed to 2000 μmol(photons) m-2s-1 for 24 hours (4).

However, other studies have shown that cyanobacterial growth can be maintained or even enhanced at high light intensities, depending on the acclimation and adaptation mechanisms of the cyanobacteria. For instance, a study by Silveira et al. (2019) found that the cyanobacterium Microcystis wesenbergii showed an increased growth rate and toxin production when exposed to 1000 μmol(photons) m-2s-1 for 12 days (5).

The response of cyanobacterial growth to high light intensity may also depend on the availability of nutrients, such as nitrogen and phosphorus, which are essential for photosynthesis and cellular metabolism. Some studies have suggested that nutrient limitation can exacerbate the negative effects of high light intensity on cyanobacterial growth, while nutrient enrichment can mitigate or reverse them (5).

Therefore, the answer to your question may vary depending on the specific conditions and parameters of your experiment. However, based on the available literature, it seems possible that the growth of Synechocystis sp. PCC 6803 can be completely halted at very high light intensities, such as those exceeding 1000 μmol(photons) m-2s-1, especially if the exposure period is long or repeated, and if the nutrient supply is low or limiting.

(1) Light intensity and spectral distribution affect chytrid infection of .... https://www.cambridge.org/core/journals/parasitology/article/light-intensity-and-spectral-distribution-affect-chytrid-infection-of-cyanobacteria-via-modulation-of-host-fitness/D491C209ABF241D5A5D28DBDBDFE0FA9.

(2) Enhancing photosynthesis at high light levels by adaptive ... - Nature. https://www.nature.com/articles/s41477-021-00904-2.

(3) Effects of Light Intensity and Exposure Period on the Growth and Stress .... https://www.mdpi.com/2073-4441/12/2/407.

(4) Mutations in hik26 and slr1916 lead to high-light stress tolerance in .... https://www.nature.com/articles/s42003-021-01875-y.

(5) Effects of light intensity and nutrients (N and P) on growth, toxin .... https://link.springer.com/article/10.1007/s10750-021-04649-z.

DOSAGE DE CHLOROPHYLE PAR SPECTROPHOTOMETRE

1) Filtration

L’eau doit être filtrée le plus rapidement possible après le prélèvement par WHATMAN GF/C

(porosité : 1,2μm).

 Placer une membrane sur le support ;

 Appliquer le vide et filtrer l’Ech. en prenant soin de l’agiter ;

 Laisser fluer l’air quelques instants ;

 Mettre le filtre dans le tube prévu à cet usage et si possible commencer l’extraction ;

 Placer aussitôt le filtre à l’abri de la lumière dans une feuille d’aluminium.

2) Extraction des pigments

 Introduire le filtre dans un tube à centrifuger et ajouter 10 ml d’acétone à 90% ;

 Déchiqueter le filtre à l’aide d’une baguette, boucher et ajouter ;

 Laisser l’extraction se poursuivre, 20h au réfrigérateur dans l’acétone à 90% ;

 Laisser revenir à température ambiante ;

 Centrifuger 1min, retirer les tubes et agiter légèrement ;

 Centrifuger à nouveau 5 à 10 min à 3000-4000 tr.min-1 ; les tubes doivent rester bouchés.

3) Mesures d’absorbance selon la méthode de Lorenzen

 Transférer le surnageant dans la cuve à l’aide d’une seringue de verre ;

 Mesurer les absorbances brutes des extrais non acidifiés (𝐴𝑏665

𝑛𝑎 et 𝐴𝑏750

𝑛𝑎 ) (𝐴𝑏750

𝑛𝑎 < 0,005) ;

 Acidifier par l’addition d’une goutte d’ac. chlorhydrique (0,3 mol.l-1), attendre 2 à 3 min et

mesurer les absorbances (𝐴𝑏665

𝑎 et 𝐴𝑏750

𝑎 ).

 Le blanc de cuve : remplir les deux cuves avec l’acétone 90% ; mesurer les absorbances

(bc750 , bc665) ;

 Mesurer le blanc de turbidité sur chaque Ech. à λ=750 nm ; (Ab750 - bc750).

4) Calcules, Expression des résultats

 Avant l’acidification : 𝐴665

𝑛𝑎 = (𝐴𝑏665

𝑛𝑎 – bc665) – (𝐴𝑏750

𝑛𝑎 – bc750).

 Après l’acidification : 𝐴665

𝑎 = (𝐴𝑏665

𝑎 – bc665) – (𝐴𝑏750

𝑎 – bc750).

 Les concentrations de chlorophylle a et de phéopigments a :

[Chlorophylle a] (mg.m-3) =

26,7(𝐴665

𝑛𝑎 −𝐴665

𝑛𝑎 )∗𝑣

𝑉∗𝑙

[Phéopigments a] (mg.m-3) = 26,7(1,7𝐴665

𝑛𝑎 −𝐴665

𝑛𝑎 )∗𝑣

𝑉 ∗𝑙

V : volume d’eau filtrée (litres)

v : volume de solvant d’extraction (millilitres)

l : longueur du trajet optique de la cuve de mesure (centimètre)

To control green microalgae contamination in Blue-green algae cultures adding Dichlorvos chemical could be a chance.

Adherence of bacteria in general, and cyanobacteria specifically, is a common phenomenon, often enhanced by stress conditions such as starvation or cyanobacteria access light. Temperature increase could also result in this phenomenon. Maybe they are lacking some nutrients?

I wouldnt go for BG-11. It is a nutrient reach medium. Instead go for a minimal nutrient medium and grow in light. Then, during exponential phase you will obtain more cyanobacteria on the expense of other not light harvesting microorganisms.

But ... You will need more steps. The most common is dilution protocols and growth on mixture of antibiotics which kill bacteria but cyanobacteria are somewhat resilient (to some extent) to them. For example see this paper:

You dont have to have axenic cultures to perform a pcr. You should use genus specific genes of interest in order to perform pcr for identification (barcoding) your species. In the case of cyanobacteria and algae, use rbcL gene (large subunit of Rubisco).

If you would like to isolate your algae/cyanobacteria, use minimal medium and grow in light. During exponential phase of your algae/cyano, you would be able to assume that these are the main constituents of the consortia, as the other organisms will have difficulties to grow.

Hi Seridi,

Cylindrospermopsis is a heterocyte-forming genus. So, if this genus is the focus of your study and you do not care about other non heterocytous genera (e.g. Oscillatoria, Phormidium, Leptolyngbya, etc), I would recommend BG110 (Same recipe as BG11 but without nitrogen source). In this way you will get rid of many genera that do not fix nitrogen and will make the isolation process a bit easier.

When it comes to isolation, I recommend this book chapter to you:

Rippka R. Isolation and purification of cyanobacteria. Methods Enzymol. 1988;167:3-27. doi: 10.1016/0076-6879(88)67004-2.

Briefly, the basic idea is to spread your crude material on the plate, incubate, then choose single filaments and transfer those on fresh plates. Depending on your sample and experience, it may take multiple transfers until you manage to get unicyanobacterial cultures so you need to be very patient and keep trying until you succeed!

best of luck with your experiments!

Maria

Thx Naresh for your kind answer.

What would interest me most would be measurements of the methane production of cyanomats, as they belong to the most productive systems we know of in the range of rainforest.

We observe now how they start to take over many systems from rivers to lakes up to whole coastal areas where they are spreading now.

Just some examples - as it is becoming a global problem...

Thus their methane emissions could become substantial on a global scale.

All the best

Jan

But the plane of division of the cells in the image you have sent me is rather horizontal than vertical, one might expect the latter if lateral protuberances are to be formed out of a division to eventuate in branches. These could also be cases of sporadic biseriate arrangement of cells in Stogonema filament which is not unheard of.

We are using cycloheximide at the concentration of 500 mg in one liter of the medium since 1973. It will inhibit the growth of many fast growing .fungi. However, dermatophytes are resistant to cycloheximide.

You need to keep in mind that all fossil ”cyanobacteria” species are morphological species, not phylogenetically distinctive species. in this sense, these calcimicrobe species are comparable to ichnospecies. We are not even sure whether they are really remains of cyanobacteria or not. You may look for the definition of “calcimicrobe” (James and Gravestock, 1981) that this just indicates calcified microfossils.

Among these calcimicrobes, only Girvanella and Hedstroemia (and related morphogroups) can be relatively reliably classified as cyanobacterial fossils. Other famous calcimicrobes, such as Epiphyton or Renalcis, are often classified as Microproblematica.

Kristova Yubilius Indrataruna I have two suggestions for you: 1) Use ASNIII medium for marine CB: BG-11 was elaborated for freshwater CB; 2) Use very purified agarose (like Kim Sea) instead of either agar or gellan gum because very purified agarose does not support the proliferation of heterotrophs. Good luck. Igor Brown

Hello, Rosalina, how are you? I'm not sure how to cultivate just a single species, but a cultivation technique that has worked well in our laboratory is to prepare a bacterial culture of raw rice with tap water... just break up a few pieces of rice grain, add to a falcon tube, put about 10mL of water in the tube, close it and wait about 2 days for there to be a considerable amount of bacteria, then just use that water in isolated cultures of protists to feed them whenever you need to. I hope to help a little with my comment, hugs!

You might try KOH for Microcystis and Woronichinia, but that will kill the cells. However, it will take at least 24 hours for the cells to lyse. I don’t know of any method for breaking up filaments.

When dilutions work but the neat/pure sample don't work the causes are most likely to be matrix effects or user error. Dilution decreases concentration of interfering substances in the matrix. Depedent on your isolation method there may be something in your solution that is interfering with the assay. High detergent and alchohol concentration are common examples of this.

Examples of methods to resolve this would be some chromotography approaches(immobilize on column and exchange buffer) or much cheaper dialysis into another buffer. However, it would be suprirsing to see such a dramatic matrix effect with zero change in signal without this always occuring when you run the test. Is this the first time this assay was run by your group?

Second just to confirm the pure sample had no change in signal from your 0 ng sample?

Formally, "uncultured Cyanobacterium sp. clone" should means that your hit is an uncultivated bacterium belonging to the particular genus, Cyanobacterium (Oscillatoriophyceae:Chroococales:Geminicystaceae). However, "sp." could be added by mistake, and then it could mean any cyanobacteria (e.g., Nostocales). For verification, it is better to blast your closest hits or exclude uncultured sequences from blast. But do not limit your search to type strains - their list among Cyanobacteria is far from complete.

I think only old edition books are available

📷

I THINK YOU SHOULD SEE NEEDHAM J G AND NEEDHAM'S ""FRESHWATER BIOLOGY"" BOOK AND ONE MORE Ponds-Small-Lakes-Microorganisms-Naturalists BY Brian-Moss. THESE HELPED ME A LOT IN MY RESEARCH OF ALGAE.

Thank you very much Ana T Castro-Castellon , for this good advice.

So far I haven't tried BG11, I have cultured using f2 media with modified NP ratio with aeration, next I will try BG11. Then, for light conditions using a standard lamp but the lux has not been measured, is there any recommendation on how many lux of light is needed, as well as other parameters such as temperature?

Thanks

thank you. one last question. what will be a fit substrate for soil protease? there are a number of options(if you know)

The magnification is too low to give a positive identification. Judging from the size and coulour I think it could be Tetraedron minimum or Chlorotetraedron incus. I do not think it is P. kamillae, because the colour does not match the one of Xanthophyceae, also the shape is slightly different, P. kamillae is more 'rounded'.

Stromatolites are layered biochemical accretionary structures formed in shallow water by the trapping, binding and cementation of sedimentary grains in biofilms (specifically microbial mats), especially cyanobacteria. They exhibit a variety of forms and structures, or morphologies, including conical, stratiform, domal, columnar, and branching types. @Bachir Achour

Try this!

We have used an antibiotic method to produce axenic alga cultures. This might work with your cyanobacterium. If you talk to a microbiologist, they have a kit used to test sensitivity of bacteria to many antibiotics. Your culture is spread on agar and different paper disks containing different antibiotics are placed on the agar plate. Look for a clear zone around a paper disk where the bacteria are killed but your cyanobacterium is not affected. This tells you which antibiotic to add to your culture medium. Our technique was published previously as “One step antibiotic disk method for obtaining axenic cultures of multicellular marine algae.” Bradley et al. 1988. Plant cell, tissue and organ culture 12: 55-60. Available on Peter Bradley’s ResearchGate place. Good luck with your work.

This looks like Gloeotila pelagica, a green alga.

After more than two years, it is time to solve the puzzle: we learned that our strain did not grow in BG-11 but was perfectly fine in Z-medium. Why? I don't know. But once we switched media, everything was fine.

Why do you want to align multiple genomes? What is your aim?

Ask this question to your course leader and understand the concepts of your the analysis you want to perform. Those tool you mention and the tool recommended in above post are not meant to align genomes.

Useless processing without proper understanding of aim would not lead you anywhere.

Fucus has been used for hundred of years by humans and in agriculture.

Cyanobacterial symbioses are no longer regarded as mere oddities but as important components of the biosphere, occurring both in terrestrial and aquatic habitats worldwide. It is becoming apparent that they can enter into symbiosis with a wider variety of organisms than hitherto known, and there are many more still to be discovered, particularly in marine environments. The chapters cover cyanobacterial symbioses with plants (algae, bryophytes, Azolla, cycads, Gunnera), cyanobacterial symbioses in marine environments, lichens, Nostoc-Geosiphon (a fungus closely related to arbuscular mycorrhiza fungi) symbiosis, and artificial associations of cyanobacteria with economically important plants. In addition, cyanobiont diversity, sensing-signalling, and evolutionary aspects of the symbiosis are dealt with. Renowned experts actively involved in research on cyanobacterial symbioses deal with ecological, physiological, biochemical, molecular, and applied aspects of all known cyanobacterial symbioses. This volume on cyanobacteria in symbiosis complements the two earlier volumes on cyanobacteria published by Kluwer (Molecular Biology of Cyanobacteria, edited by D.A. Bryant and Ecology of Cyanobacteria, edited by B.A. Whitton and M. Potts). Together, the three volumes provide the most comprehensive treatment of cyanobacterial literature as a whole. The book will serve as a valuable reference work and text for teaching and research in the field of plant-microbe interactions and nitrogen fixation.

Omnia Rabea The structure of Cynabacteria

This one is my personal favourite Climate Change and Cynabacteria

Hello Dear, these articles may help you:

repository deposition of the raw proteomics data is required for most journal as a way of validating the authenticity of the data in the manuscript. It will also increase the global publicity of your manuscript

Thanks Anju Kaushal for nice suggestion.

@sezgi adalioglu thanks for you response it give more insight to understand about the condition of chlorella sorokiniana. Please is there any documents I can read to know about this? If they is any I need your help to send it for me. Thank you

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 (

Dear Maria,

Hereby, please find two recent publications about cyanobacteria, which may help you find what you want to know about its methodology.

The short communication published in 2021 is a good one to refer to its references and gives a good idea of overall researches around Cyanobacteria.

The paper published in 2019 may give you tips in methodology of Cyanobactetia grown in harsh environmental situations.

Good luck.

You may add some growth inducers to your culture media.

Hi Lana Ezzdeen

I would suggest that you use the single cell (or filament) technique to start, in liquid media. This medium can be BG-11, ASM-1, WC medium, among other options. I mostly used WC medium for freshwater species (Guillard RRL, Lorenzen CJ (1972) Yellow-green algae with chlorophyllide C 2. J Phycol 8:10–14 doi:https://doi.org/10.1111/j.1529-8817.1972.tb03995.x).

This is the way I used to do it:

I hope that those tips help you isolate your organisms!

All the best,

Guilherme

*We used hydrophilic cotton on the bottom of the pipette to prevent contamination, and also adapted a silicon tube that we would connect to the pipette and would suck with our mouths, with hydrophilic cotton on the "mouth" end. Sounds like it doesn't work, but it is super effective and we didn't have problems with contamination, not even with bacterial contamination when working with axenic cultures.

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!

Dear Niharika,

I hope the Antioxydant protocols found in these 2 articles (attached) will help you in your research.

Best wishes