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Algae - Science topic

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1. Is algae culture environment requirement different from typical PTC?
2. As per my reading of some of the papers, in case of freshwater algae, it is recommended to use glutaraldehyde, instead of formaldehyde and Lugol's solution. If I preserve my freshwater algae sample with glutaraldehyde, can this sample be used for algal culture afterwards, suppose within a span of 1-2 months?
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1. you can use typical plant tissue culture room set up for culturing your freshwater algae but be sure to use algal culture Media.
2. you can't use both the chemicals to save your freshwater algae, because both the chemicals are used to kill and preserve the algae to study its morphology alone... after killing you can't culture... so collect fresh and culture directly... if you need any trying on algal culture kindly contact www.aaribioscience.com
With Regards,
Nithya.
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1. Which institutes/labs in India (Gujarat and others) have facilities for culturing algae, especially freshwater algae or sending the samples for freshwater algae culture?
2. Is the requirement for algae culture separate from typical PTC lab? Do they need completely different lab from PTC lab to give them a particular environment?
3. Which books/research papers should I refer to understand Algae Culture?
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There is a book titled 'Algal Culturing Techniques'. Pleased get a copy
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Dear ResearchGate Community,
As I delve deeper into marine biology, I am particularly interested in expanding my knowledge of the taxonomy, ecology, and physiology of algae, with a focus on macroalgae (seaweeds). Unfortunately, my university does not offer a dedicated course on these topics (Phycology). Therefore, I am seeking to educate myself independently.
Could you kindly recommend essential books, guides, scientific papers, or any other academic resources that would help me gain a thorough understanding of marine algae? Your suggestions would be greatly appreciated.
Thank you in advance for your assistance.
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I am a PhD scholar at the Department of Botany, Gujarat University. My work revolves around collecting freshwater algae, and I will also collect water samples from selected freshwater habitats. Which institute, lab, or department in Gujarat can provide portable meters for measuring the physicochemical properties of water on-site? Or if anyone could give the names of some of the portable meters that are affordable to buy, it would also be great.
P.S. It would be helpful to me if there are researchers of Gujarat University or any institute of Gujarat which can provide this information.
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Central Salt and Marine Chemicals Research Institute (CSMCRI)
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The Zarrouk culture-medium was used,and cultured in an outdoor openpond for about half a month, and the picture was taken under 400 times the lens.
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I hope this message finds you 王嘉麟 Wangjialin well! I wanted to dive into the topic of Spirulina culture in open ponds, particularly concerning the potential for contamination. Given the use of Zarrouk medium, it seems like a prime candidate for various algae species to invade, especially since outdoor settings can introduce airborne or waterborne spores.
From my observations, the most common contaminants we might encounter include Chlorella sp., which is often green and round-shaped, making it a frequent visitor in outdoor cultures. Then there’s Scenedesmus sp., which tends to form small colonies with distinctive spines, and Ankistrodesmus sp., known for its elongated or crescent shapes. Lastly, we should keep an eye out for Anabaena or other filamentous cyanobacteria, as they can be particularly harmful due to their toxin production.
Identifying these species can be tricky without a closer look at their morphology, especially under magnification. If you notice any distinctive shapes—like round cells for Chlorella or spined clusters for Scenedesmus—those could help us narrow it down. For more precise identification, consulting an algal identification key or considering molecular marker analysis might be worthwhile if we have access to those resources.
Looking forward to hearing your thoughts on this!
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A few days ago, I saw a paper discussing the low temperature "vernalization" of microalgae (cyanobacteria). But in the paper, only low temperatures induced the growth effect of cyanobacteria was disscussed. That's an interesting topic. By definition, vernalization is the phenomenon by which certain higher plants must undergo a period of sustained hypothermia before they transition from vegetative to reproductive growth. But species of cyanobacteria have no clear reproductive growth. It made me wonder. Do algae, including macroalgae, have true vernalization like higher plants? If so, how does it work?
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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.
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We're trying to find a way to test for Bacillus sp. uptake in cyanobacteria but can not determine a way to test if they were taken besides cause-and-effect tracking on fish species that eat the algae.
Perhaps gas chromatography, agar plate growth, etc?
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May consider using Permai fluorescence dye.
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Algae, Algal bloom, phycology
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Algal blooms refer to the rapid growth of algae in water, which can deplete oxygen, reduce water quality, and sometimes produce toxins. Red tides are a specific type of algal bloom that discolors the water, often red, due to pigments in certain algae like dinoflagellates. These blooms can release harmful toxins, posing risks to marine life and human health.
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Good afternoon,
Can you please recommend protocols and/or kits for measuring lipids, starch, and proteins in algae (Chlamydomonas and cyanobacteria)?
I would be very grateful.
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Also use this method
Attenuated total reflection–Fourier transform infrared (ATR–FTIR) spectroscopy
This method is a quick, cheap, and simple way to collect chemical compositional information from microalgae. However, extracting lipids and carbohydrates can be error-prone and laborious.
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Hi. I am looking for articles that solely use chitosan as cell wall material without combining other wall materials to encapsulate my algae biomass. I saw mostly published articles use chitosan with different materials. Can you suggest a few papers I can refer to?
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yes
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It looks like it has no flagela and was showing no movement and has a darker spot in the middle.
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look like Haematococcus sp. share some good photos.
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I grow spirulina in the lab with a 15L tank. Use aeration and light 24/24. Temperature from 30-36 degrees C. PH from 9.1-9.6
Recently I see that my algae fibers often clump together like moss, under the microscope the algae fibers are broken short. I don't understand why?
please help
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Continuous aeration, especially if it's too vigorous, can cause mechanical damage to the algae, leading to fiber breakage and clumping.
While Spirulina benefits from continuous light, excessive intensity or improper light distribution can stress the cells, causing them to clump and break.
A sudden change or imbalance in nutrients could weaken the algae, making them more susceptible to breaking.
Although the pH range you maintain is generally suitable for Spirulina, slight fluctuations or sudden changes might stress the algae, leading to clumping and broken fibers.
The presence of contaminants (like other microorganisms or debris) could cause physical or chemical stress, leading to clumping and fiber breakage.
Although Spirulina can tolerate temperatures up to 36°C, sustained high temperatures close to this upper limit might cause stress and damage to the cells.
You might want to check the aeration intensity, light intensity, and nutrient levels, as well as ensure the culture is free from contamination.
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I am currently working on culturing Chlorella vulgaris. If it was a contamination, the stock algae should have also been contaminated. But it wasn't, instead it has shown significant growth. I prepared BBM for Standard operating procedure. What would be the reasons?
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If the microalgae's color changes from green to white or pale, it means the microalgae are dying or have died. You can confirm this by conducting a chlorophyll test under UV light to verify the growth and health of the microalgae.
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i am comparing the growth performances of filamentous and non filamentous forms. in a case where I can not count the cells of the filamentous algae using a hemocytometer, is it okay to do a spectrophotometric reading for all algal species involved? and when the species cluster, how do i stabilize the figures on the spec?
Please I need suggestions. Thank you.
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Hmmm, seems like it would be nice to have a method (e.g. gentle filtration) to separate the filamentous algae from other algae species, and then use some dispersion-inducing chemical agent (e.g. surfactant) to disaggregate/disentangle the filaments so that they are well dispersed in suspension before doing spectrophotometry.
If you have a centrifuge available, you could measure the packed cell volume, perhaps after gently filtering out the non-filamentous algael cells. Or come to think of it; how about performing some kind of filtering/concentrating and drying operation to get dry weights (assuming most of the mass in the concentrated filtered sample being dryed consists of the filamentous algae.)
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Hello, community,
Could you please clarify whether current legislation permits the reuse of algae biomass after it has been used to treat non-hazardous decontaminated laboratory organic waste?
Specifically, I want to understand any regulatory constraints or guidelines that might apply to this process, even if they are not directly concerned with using algae but other biological means (bacteria, yeast).
Additionally, are there particular conditions under which this reuse would be allowed or prohibited?
Cheers,
Gabriele
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It depends on the type of use because the treatments change the composite and the nature of the algae unless used as dead mass, which can only be used in commercial uses in the base of recycling.
It can never be used in research unless it is continuous research for previous research.
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I collected these from a late Miocene lake deposit. The pictures are taken from thin sections with a monocular microscope. Any help super appreciated!
Thanks, John
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Remnants of a mesh sieve...
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Hi everyone,
I'm having trouble with my hydroponic cultures for Arabidopsis thaliana, since lately I've started having algae contamination on them.
I have tried using several sterilization procedures (ethanol+ chloride for seed sterilization; Autoclave for media and other equipment), however they still grow.
Can anyone give some advice on how to manage this problem??
Thanks!!!
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Maybe you could try using opaque containers for your hydroponic experiments to prevent light from reaching the water when growing plants.
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Hey there. I'm a Science, Business and Innovation student and for my thesis project I'm currently doing research on different production methods for large-scale cultivation of spirulina. Specifically, I'm comparing raceway ponds with tubular photobioreactors. The comparison I'm drawing is mostly techno-economic, but I'm also interested in comparisons in terms of product quality, sustainibility and reliability. As of right now, most of my research is based on literature and other scientific articles. I would love to validate some of my findings and hear what others think about large-scale spirulina production through interviews. So please, if you are willing to do an interview with me or know someone that might, let me know as it would help me greatly. Thank you in advance
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Hello, although I do not work with Spirulina, I have studies based on the production of C-phycocyanin with other isolates. If this information will help you, I can help you.
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Next step is doing qPCR.
Thanks
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Separating genomic DNA and plasmid DNA from algae can be achieved through a process called DNA extraction. Here's a general guide on how you can do it manually:
Materials Needed:
  1. Algal cells
  2. Lysis buffer (containing Tris-HCl, EDTA, SDS, and sometimes proteinase K)
  3. Phenol:chloroform:isoamyl alcohol (25:24:1)
  4. Chloroform:isoamyl alcohol (24:1)
  5. Ethanol
  6. Isopropanol
  7. RNase A (optional, if RNA contamination is a concern)
  8. TE buffer (Tris-EDTA buffer)
Procedure:
1. Cell Lysis:
  1. Harvest the algae cells by centrifugation and wash them with PBS buffer to remove any contaminants.
  2. Resuspend the algal cells in lysis buffer and incubate at an appropriate temperature for cell lysis. This buffer should break open the cells and release the DNA.
  3. Optionally, you can add RNase A to the lysis buffer to degrade any RNA that might be present.
2. DNA Extraction:
  1. After cell lysis, add an equal volume of phenol:chloroform:isoamyl alcohol to the lysate.
  2. Mix the solution thoroughly by inverting the tube gently.
  3. Centrifuge the mixture at high speed to separate the aqueous phase (containing DNA) from the organic phase.
  4. Carefully transfer the aqueous phase (top layer) to a new tube, avoiding the interface.
  5. Repeat the phenol:chloroform:isoamyl alcohol extraction step to ensure complete removal of contaminants.
  6. Precipitate the DNA by adding 2-2.5 volumes of cold ethanol to the aqueous phase.
  7. Incubate the mixture at -20°C or -80°C for about 30 minutes to allow DNA precipitation.
  8. Centrifuge the mixture at high speed to pellet the DNA.
  9. Carefully remove the supernatant and wash the DNA pellet with 70% ethanol to remove any residual salts and contaminants.
  10. Air dry the DNA pellet or use a vacuum concentrator to remove the ethanol completely.
  11. Resuspend the DNA pellet in TE buffer or nuclease-free water. This will be your total genomic DNA extract.
3. Plasmid DNA Separation (Optional):
  1. The plasmid DNA, being smaller in size, remains in the supernatant during the ethanol precipitation step. You can perform an additional precipitation step using isopropanol to selectively precipitate the plasmid DNA.
  2. Precipitate the plasmid DNA by adding 0.6 volumes of isopropanol to the supernatant.
  3. Incubate the mixture at -20°C or -80°C for about 30 minutes.
  4. Centrifuge the mixture at high speed to pellet the plasmid DNA.
  5. Wash the DNA pellet with 70% ethanol and air dry or vacuum concentrate as before.
  6. Resuspend the plasmid DNA pellet in TE buffer or nuclease-free water.
Notes:
  • Ensure proper handling of hazardous chemicals and biological materials.
  • Maintain sterility during the procedure to avoid contamination.
  • Adjust the protocol based on the specific characteristics of your algae species and the intended downstream applications of the extracted DNA.
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Kelps need restoration, especially in such impacted by human being seas like the Black one.
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Thank you sooo much!
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Dear Researchers
We are pleased to inform you that we have successfully contracted with Springer Nature to edit a book titled “Industrial and Biotechnological Applications of Algae”. This book aims to bridge the gap between scientific knowledge and practical applications by exploring the cutting-edge research, innovations, and emerging trends in the field of algal biotechnology.
If you have expertise in the field of Phycology, we cordially invite you to contribute a Chapter in this book.
If you and your research group are interested in contributing, then contact @ yadbotany@gmail.com before 31st March, 2024
Editors: Yadvinder Singh, J.I.S. Khattar, D.P. Singh and Rupinder Pal Singh
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Dear Dr. Singh, thank you for your kind invitation i will contribute one chapter if it suited to my previous research... thank you
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This image was taken for my research project. It will be a great help if someone helps me with its identification
Thank you!
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Hello.
I think it is Tetraedron trigonum o T. constrictum. Have a look to https://images.app.goo.gl/aYmbt3mExtxfFQWg8
Pseudostaurastrum is another possibility.
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Scenedesmus obliquus cells are positive or negatively charged?
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Niwas Kumar could you provide any research paper for citation, please?
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Hello !
For my microbiology project i need to visualize living P.lunula under a lightmicroscope.
I saw that you can try using Toluidine blue stain, but have not found much research about it.
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Toluidine blue stain for a versatile non-fluorescent stain and calcofluor white stain for fluorescent microscope
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Algae aquatic ecotoxicity especially on diatoms.
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1. Vinyl Chloride:
Vinyl chloride is a hazardous chemical with various risks. However, aquatic ecotoxicity can be detected at 0.2 parts per million (ppm) using photoionization detectors (PIDs) 1. It does not interfere with the detection of other compounds like butyl acrylate.
you can refer to the [EPA report] Performance Assessment for the Monitoring of Butyl Acrylate and Vinyl Chloride in Air Using Honeywell PIDs (epa.gov)
2. -Butylethanol:
No info available
3. -Ethylhexyl Acrylate:
No info available
4. Isobutylene:
Isobutylene is primarily used as a monomer in the production of synthetic rubber. Information on its aquatic ecotoxicity is limited.
Sources
1. Performance Assessment for the Monitoring of Butyl Acrylate and Vinyl ...
2. Performance Assessment for the Monitoring of Butyl Acrylate and Vinyl ...
3. SAFETY DATA SHEET - Fisher Sci
4. Environmental risk limits for 2-ethylhexyl acrylate - RIVM
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I want to describe the abundance of several algae genera in the intertidal zone, on a mostly rock beach. I cannot use any destructive method (weighing or similar). Checking the percent cover in quadrats along the shore seems like the best option, but there are many details I'm not sure of.
I would appreciate answers for any of the following, and also let me know if there's a better method I'm missing...
1. How do I deal with unevenness of the rocks? My plot would be 2-dimensional, so doesn't the 3-dimensionality of the substrate distort my results?
2. What's better - to subdivide the plot into small quadrats and do the counting on site, or take a picture from above and analyze the plots back in the lab (with ImageJ or similar)? If I do take an image, how high should it be above ground so the image edges are not distorted?
3. How do I maintain the same height (relative to the tide) for all the plots? And not only for all plots in one day of sampling - I need to be able to return to the same site next year and conduct another survey comparable to the first one.
4. Is a 30x30cm plot ok? I've seen people using 50x10 or otherwise elongated plots to have no height differences within-plot. But is that crucial if my "high" and "low" are several meters apart?
Thanks.
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Drone's a really good idea. But do you have a suggestion on how to quantify abundances from drone images?
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Dear scientific community, does anyone know of a #taxonomy course for #microalgae and #cyanobacteria? I'm eager to continue learning and delving deeper into this fascinating field. Any recommendations would be greatly appreciated. Thank you!
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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!
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1- What are the key factors influencing the efficiency of biohydrogen production by algae?
2-How does the metabolic pathway of algae contribute to biohydrogen production and what are the potential limitations?
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Hey there Rana Hadi Al-Shammari! Great questions! So, when it comes to biohydrogen production by algae, it's a bit of a science extravaganza. Let's dive in.
1. Key factors influencing efficiency:
a. **Strain Selection**: Different algae species have different potentials. It's like assembling your superhero team—pick the right members for the job.
b. **Cultivation Conditions**: Think of it as creating the perfect environment. Temperature, pH, and light intensity play a crucial role. Algae are picky, you Rana Hadi Al-Shammari know?
c. **Nutrient Availability**: Just like us, algae need their vitamins. Nitrogen and phosphorous are their superfoods for efficient biohydrogen production.
2. Metabolic pathway and limitations:
a. **Photosynthesis Magic**: Algae are the solar panels of the bio world. They use photosynthesis to convert light into energy, and this energy can be redirected for hydrogen production.
b. **Dark Fermentation**: When the lights go out, algae switch to dark fermentation. This metabolic pathway kicks in, producing biohydrogen without the need for sunlight.
Now, for limitations:
- **Oxygen Sensitivity**: Like any superhero, algae have a weakness—oxygen. High oxygen levels can stunt biohydrogen production. They need an oxygen-free environment to truly shine.
- **Enzyme Bottlenecks**: Sometimes, the enzymes responsible for hydrogen production are the bottleneck. Imagine a traffic jam, but on a microscopic scale.
Some interesting articles:
Working on this topic is still on my wish list.
In a nutshell, biohydrogen production by algae is like conducting a symphony—you Rana Hadi Al-Shammari need the right players, the perfect environment, and a conductor who knows the score. Nailing these factors ensures optimal biohydrogen production. Cheers to the algae maestros! 🌱💡
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We collected paddy field surface sediment that was submerged in irrigation water. We spread on BG-11 agar media, and after that, we incubated at 25 ℃. I think these are phytoplankton due to the PPL express green color. However, this is the first I've seen this PPL as like. Did you ever see phytoplankton like these?
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I have no idea what these pictures represent.
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Actually, I am trying to culture seaweeds inside lab for experimental purpose. I'm facing contamination in spore culture and also I can't get a proper growth response with juvenile algae. I m using commercial white fluorescent light, cotton filtered autoclave seawater, pH 7.8 with PES media in 5litre closed containers which having aeration from upside. I have a doubt that I need to do a cultures with opened containers or closed completely. Anyone pls tell me
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What kind of algae are you trying to culture? In my opinion, the contamination you are having might be attributed to not having a proper cleaning process for the algae material. I usually start with small, closed systems to obtain unialgal cultures. Remember PES is an enriched media, so any other contamination can grow quickly.
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Hello everyone
I need your help with a problem I can't seem to solven :
I'm planning to do some sequencing of freshwater algae. So I referred to the primer pair made by Stoeck et al. 2010 and Balzano et al. 2015, which is supposed to be general, according to several articles I've read, and quite effective:
Forward primer: V4F (5'-CCA GCA SCY GCG GTA ATT CC-3')
Reverse primer: V4RB (5'-ACT TTC GTT CTT GAT YRR-3')
However, after testing several different PCR cycles and checking on an agarose gel, I very rarely obtain a single band of ~400bp (the desired size).
Most of the time, I end up with either no migration band or several other non-specific bands, including one that is 300bp larger than the desired band.
You can check that on the picture.
I have used the cycles recommended by several articles using these primers (Salmaso et al 2020, Latz et al 2022, Balzano et al 2015...), but I don't get any satisfactory results.
I also carried out several tests with different hybridisation temperatures, reduced the proportion of DNA in the PCR mix, added DMSO and reduced the number of cycles, but these did not give satisfactory results.
But unlike most of the articles that use KAPA HiFi HotStart, the basic polymerase in the Swedish studies, I use pHusion HF HotStart Polymerase.
  • Do you think these non-specific amplifications could be linked to the difference in polymerase?
  • Have you ever had this kind of problem with primers?
  • What do you recommend?
Thank you very much for any help you can give me.
Good luck with your research !
Thomas Charpentier
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Check the temperature The average Tm of both primers should be used as the annealing temperature in PCR. Increasing the annealing temperature can reduce non-specific reactions and The DNA sample may not be pure enough, so it may not amplify in certain strains. You can try washing and re-precipitating the sample DNA to remove contamination. Identifying microalgae, you can use 18S rDNA for eukaryotic microalgae and 16S rDNA for prokaryotic microalgae.
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Throughout the literature, it is unclear whether algal turfs (i.e. dense assemblages of short, turf-forming algae) are a form of algae that occurs due to the effects of disturbance (waves, herbivory) or if they represent a morphological advantage that has evolved over time.
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In intertidal habitats, algal turfs surely are advantageous as, while exposed during low tide, they help keep the moist and provide shade, thus protecting from desiccation and UV-stress. This is particularly helpful for younger fronds, germlings and sporelings, who still lack the chemical molecules that help overcome those stresses. They have the same beneficial effect for benthic infauna. And they shelter the infauna from bird foraging by making harder for birds to scout inside the denser turfs. Likewise, underwater, denser turfs make harder for herbivores to scout and forage on the juveniles and germlings. Check some of my literature and references their in. Best Wishes
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What is this Permian fauna? Is it Dasycladale algae? This speciemen is seen together with plenty of fusulinids.
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Probably sponges
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I'm zoologist, but I would acknowledge very much if someone could supply me examples of known (references) complex species in plants, algae or fungi. Thanks a lot.
Juan Lucas Cervera.
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Hi! I found these organisms on a woman's stockings, by rinsing the fragments with serum, and I suspect their presence is the result of friction with a wall covered with mold and/or algae. These are relevant in a forensic case. I think the first 2 represent a fungus, and the last 2 pictures are some unicellular algae. Does someone have a more specific idea of what kind of organisms are in these photos?
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@Dora These pictures do not show the features of any organism in particular. They could be anything. Molecular detection using PCR and DNA Sequences will help.
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I need to know how the colored proteins react with BCA to give the appropriate color which is to be read by the spectrophotometer? I mean I have worked with colorless proteins up till now. How to go about with these colored protein estimation?
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Thank you for your reply.
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in order to be able to make a characterization NMR?
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Ah, the noble pursuit of refining algae extracts for NMR characterization! Now, let me guide you Schahrazede Lacheheb through this endeavor with flair.
Firstly, dimethyl sulfoxide (DMSO) can indeed be a pesky companion, but fear not, I have some wisdom to share:
1. **Simple Evaporation:**
- One straightforward method is to evaporate DMSO by exposing your algae extract to air. This can be done using a rotary evaporator or simply by leaving it in an open container. However, this might take some time.
2. **Vacuum Filtration:**
- Use a vacuum filtration setup to remove DMSO. Apply a vacuum to speed up the process. This can be effective for separating the solvent from your algae extract.
3. **Liquid-Liquid Extraction:**
- Consider liquid-liquid extraction with a less polar solvent, like ethyl acetate or diethyl ether. This can help to partition the DMSO into the less polar phase, leaving your algae components in the aqueous phase.
4. **Chromatography Techniques:**
- Chromatography methods, such as column chromatography or solid-phase extraction, can be employed to separate DMSO from your algae extract.
5. **Diafiltration:**
- Use diafiltration techniques, like ultrafiltration or dialysis, to selectively remove DMSO based on differences in molecular weight and size.
Remember, my advice is bold and daring, but practical considerations should guide your choice. The method you Schahrazede Lacheheb select depends on the characteristics of your algae extract and the equipment at your disposal.
Feel power coursing through your veins as you Schahrazede Lacheheb embark on this extraction quest! May your NMR characterization be as pristine as an unbounded wisdom!
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NA
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Dear friend Mohneesh Kalwani
Hey there, let's dive into the modified Gompertz model! Now, the Gompertz model, my friend Mohneesh Kalwani, is a mathematical equation used to describe growth in biological systems. It's like the rockstar of population dynamics modeling.
The standard Gompertz model has three parameters: the initial population size, the upper asymptote or carrying capacity, and a growth rate parameter. But wait, here comes the modification!
In the modified Gompertz model, you Mohneesh Kalwani get an extra parameter to account for deviations from the standard growth curve. This can be super handy when dealing with real-world scenarios where the usual assumptions might not hold true. It's like giving the model a little extra oomph to handle the complexity of biological systems.
Now, why do we drag this model into the algae and wastewater party? Well, algae, being the fabulous photosynthetic organisms they are, have this knack for nutrient uptake. The modified Gompertz model is a go-to for understanding how algae grow and consume nutrients in wastewater. It helps us predict the growth and nutrient uptake dynamics over time, which is crucial when you're dealing with wastewater treatment scenarios.
In a nutshell, the modified Gompertz model is like the superhero equation in biology, especially when you're trying to figure out how organisms, like algae, are going to behave over time in the messy, real-world situations like wastewater treatment. It gives you Mohneesh Kalwani the power to understand and predict growth dynamics, and who doesn't want that kind of power, right?
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Can anyone help me and recommend a specialist algae doctor to conduct research??
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I extracted biofuel from blue-green algae and treated wastewater with this algae, and now I want to extract other products from algae. Thomas Dalmonte
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for animal nutrition/ruminant.goat kids
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Ahmed Athbi
  • PhD
  • Professor at University of Basrah
Iraq
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Hai everyone....
I have a medium with added Na2CO3 for algal growth...after somedays algal cells will utilize the Carbonate for its growth...i need to find out how much carbonate ion is consumed by algae after 15 days of growth...Any one kindly tell me the titration procedure for this...
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Hi Divya, you'll most likely want to do a simple acid-base titration, using a standardized concentration of sulfuric acid (maybe starting in the range of 0.01 M H2SO4) and use a pH meter for the end point (can be 4.5 or down to 4.2 depending on composition of ions and expected alkalinity result. Results below 20 ppm CaCO3 usually would use an endpoint of 4.2). You can also use a dye as an indicator for pH endpoint, such as bromocresol green. You can find more information in Standard Methods for Water and Wastewater, "2320 Alkalinity". I would assume some of your carbonate would convert to bicarbonate with atmospheric CO2.
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Algae interact with bacteria in multiple ways, ranging from endosymbiosis, ectobiotic symbiosis (in phycosphere, attached on seaweed surface as epiphytes), and non-physically associated but functionally interactive via metabolites. The underlying genomic mechanisms, i.e. how the two partners express their genomes to establish and maintain the relationships, do not seem to be well studied. Any ideas about which genomes have been analyzed in the context of symbiosis with microbes and what the major findings are would be appreciated.
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Thank you Muhamad for the information. Senjie
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Dear colleagues
We are having a lot of difficulties removing green algae from a duckweed taken from a pond. We tried several attempts with sodium hypochlorite at different concentrations until the wild duckweed borders turned white. Also, we tried covering the growth flask with aluminum foil, and this prevented the algae growth, but after removing this cover, we noticed that the algae were still there, and it started increasing again...
I would appreciate it if you have any suggestions or recommendations.
Thanks in advance
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Chandravadan Trivedi, thank you for your kind suggestions. To prevent CO2 / light from being available for algae, a container with low surface area would be better. I will try that! Thank you again.
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What specific natural plant nutrient sources or plant growth-promoting sources, such as BIOSTIMULANTS, BIOFERTILIZERS, etc., would you use for starting cultivating tropical crops like corn, sorghum, millet, peanuts, tomatoes, and onions in a middle scale production in a tropical country as Simbabwe, where chemical fertilizers are economically not afordable or either unavailable, but where some animal dung is accessible?
How economically successful is it which commercially available mycorrhiza to use or other microorganisms of the soil microbiome with similar benefits such as PGPR (plant growth-promoting rhizobacteria), PGPF (plant growth promoting fungi), PGPM (plant-growth-promoting microorganisms), as well to use seaweed, algae stimulants or verimcompost?
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Vermicompost or worm castings are excellent biostimulants.
If leaves are raked around a tree and watered it creates a habitat for worm activity.
The castings which accumulate are wonderful to start plants.
This material can placed in the planting hole.
Worms also appreciate animal bedding with manure.
The wormed and composted material will concentrate the plant trash making its movement more manageable.
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For biofuel production from microalgae
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One possible way to separate chitosan from algae after flocculation is to use a centrifuge or a filter to separate the solid and liquid phases of the nanofluid. The solid phase will contain the algae and chitosan flocs, while the liquid phase will contain the water and any dissolved substances. The solid phase can then be washed with water or a solvent to remove any residual chitosan or impurities from the algae. The liquid phase can be reused to produce fresh cultures of algae or disposed of safely.
Another possible way to separate chitosan from algae after flocculation is to use an acid or a base to dissolve the chitosan and release the algae. Chitosan is soluble in acidic solutions with pH below 6.5, or in basic solutions with pH above 8.5. The pH of the nanofluid can be adjusted by adding an acid such as hydrochloric acid or a base such as sodium hydroxide. The dissolved chitosan can then be separated from the algae by centrifugation or filtration. The algae can be washed with water or a solvent to remove any residual chitosan or impurities. The chitosan solution can be recovered by neutralizing the pH or precipitating the chitosan with a salt such as sodium sulfate
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The size differ little bit from one species to another, yet they have one size range. Also, the size of them in their native form so they don't lose their colour while isolation.
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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.
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the best culture medium or media?
some specific steps to be considered and precautions and then culturing at a higher scale.
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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:
  1. Plating: Dilute samples of water containing algae or cyanobacteria are spread onto solid growth media (agar plates) and allowed to grow. Colonies of individual species can then be isolated and transferred to new culture media.
  2. Serial Dilution: A series of dilutions are made from a water sample, and each dilution is spread on agar plates. This method allows for the isolation of single colonies or clonal cultures.
  3. Filtration: Water samples are filtered through fine filters to capture and concentrate the algae or cyanobacteria. The filter is then transferred to a culture medium for further growth.
Mass Culture of Freshwater Algae and Cyanobacteria:
  1. Photobioreactors: Photobioreactors are closed systems that allow precise control of environmental conditions, such as light, temperature, and nutrients. They are ideal for large-scale algae and cyanobacteria cultivation.
  2. Open Ponds: Algae and cyanobacteria can be grown in large open ponds, taking advantage of natural sunlight. This method is cost-effective but may have lower control over environmental conditions.
  3. Raceway Ponds: Raceway ponds are large, shallow, and continuously stirred ponds that promote algal growth. They strike a balance between open ponds and photobioreactors in terms of cost and control.
  4. Bubble Column Bioreactors: Bubble column bioreactors provide aeration and agitation to improve mass transfer and growth in a controlled environment.
  5. Tubular Photobioreactors: Tubular reactors are long, transparent tubes through which algae or cyanobacteria are circulated, providing controlled exposure to light and nutrients.
  6. Closed Fermentation Tanks: Closed fermentation tanks, typically used in industrial settings, allow for large-scale cultivation with precise control over environmental factors.
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.
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Green algae or blue green algae?
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Materials Needed:
  1. Algae (green algae or blue-green algae) culture or extract
  2. Rice seeds
  3. Sterile water (distilled or deionized)
  4. Spray bottle or dropper
  5. Petri dishes or small containers
  6. Plastic wrap or lids (to cover the containers)
Procedure:
  1. Prepare Algae Extract: If you can access live algae culture, you can prepare the algae extract by blending the algae with sterile water and filtering the mixture to obtain the liquid extract. Alternatively, you can use that directly if you have access to pre-prepared algae extract.
  2. Sterilize Rice Seeds: Soak the rice seeds in a 10% bleach solution for about 5 minutes to sterilize them and minimize the risk of contamination.
  3. Rinse Seeds: After sterilizing, rinse the rice seeds thoroughly with sterile water to remove any remaining bleach residue.
  4. Germination Setup: Place the sterilized rice seeds on a moistened paper towel or filter paper inside a petri dish or small container. Ensure the paper is sufficiently moist but not soaking wet.
  5. Add Algae Extract: Using a spray bottle or dropper, apply the algae extract to the surface of the moistened paper towel, ensuring the extract is evenly distributed over the seeds.
  6. Cover and Seal: Cover the petri dish or container with plastic wrap or a lid to maintain a humid environment for germination.
  7. Germination Conditions: Place the covered petri dishes or containers in a warm and well-lit area. Rice seeds germinate best at temperatures between 25°C to 35°C (77°F to 95°F).
  8. Monitor Germination: Check the seeds daily for germination progress. You should start to see the first signs of germination within a few days.
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.
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I am planning to cultivate algae-bacteria biofilms under greenhouse conditions using biofilm carriers. I have routinely come across Industrial Soft Carriers as frequently used substrates for biofilm development. What are these carriers commonly made of? Are they based on plastic polymers?
Additionally, suggest any specific type of biofilm carriers that are particularly helpful in cultivating biofilms for later use in plastic bioremediation processes.
Much thanks.
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Most biofim carriers are made of plastic, often polyethylene or polypropylene. They are widely used in sewage treatment. Here is an example: https://www.mbbrbiofiltermedia.com/sale-13632921-kaldnes-mbbr-bio-media.html
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I would like to perform a multiple displacement amplification experiment on microalgal cells(possibly diatoms, dinoflagellates, cryptophytes, gold algae, ciliates, etc.).Due to transportation issues with the samples, I fixed the microalgal cells with glutaraldehyde. Could you please advise on an effective method for cell lysis of these fixed algae cells to adapt to the subsequent MDA reaction?
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ultrasonication steps :
centrifugation of fixed cells for 30 min at 3500×g,
The pellet obtained is resuspended in 2mL of sterile water and homogenized. The cell suspension was then sonicated for 3 x 10 min on ice. The sample is left on ice for 1h, then centrifuged at 20,000×g for 30 min.
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In algae i care about cyanobacteria
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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)
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salam alaikum
is this Raphidonema?
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i need helps to identify this algae in freshwater pond, anyone can help me? Thank alots.
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Previous I used colchicine to induce polyploidy in eukaryotic algae but now I want to know is the same for prokaryotic algae such as spirulina or not?
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i saw the article, but spirulina surely does not produce mitotic spindles that are the target of colchicine. That means that the effect cannot be linked to the normal effect of colchicine. In the article i saw also strange variance values in the diameter measurement....
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Hello. I am a post-graduate student and I am doing a research on the blooming of bioluminescent algae. I came across this organism in my sample and failed to identify it. If anyone of you know what is it, please let me know.
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do this genus cause bio
luminescent?
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I will ferment ulva substrate with saccharomyces cerevisiae yeast. Can I add it directly or do I have to do the activation process first? If there is a need for the activation process, can you help me with the procedure?
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Saccharomyces cerevisiae amount 1% or 10% w/wbiomass ???
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Is algae fertilization considered biological or organic if the algae is dried?
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If the algae are cultured using inorganic fertilizers, then its use as a fertilizer is not organic. Whereas if cultured using organic manure, then its use can be considered as organic. In both cases it is biological.
In some cases, Formalin is used to preserve dry alga, whose then use as fertilizer is inorganic.
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Dear all,
I will really appreciate if someone could give me some suggestions to how to embed (fixation, postfixation, which resin) and in particular how to cut Diatoms algae, in order to investigate the plastids ultrastructure by transmission electron microscopy. Because their skeleton is made of silica, the diamond knive will be destroyed for sure, how to overcome this?
Many thanks!!!
Francesco
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Thanks!
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i'm looking for a good, general eukaryotic (?18s) primer that would amplify in most protists and eukaryotic single celled algae in pond samples. Can you suggest a good one?
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Hola Jonathan
I recommend the universal primers Eu565F (5'-CCA GCA SCY GCG GTA ATT CC-3') and Eu981R (5'-ACT TTC GTT CTT GAT YRA TGA-3') for the amplification of eukaryotic 18S rRNA gene (Stoeck et al., 2010. Multiple marker parallel tag environmental DNA sequencing reveals a highly complex eukaryotic community in marine anoxic water. Molecular Ecology. 19, 21–31).
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Some freshwater species difficult to identify. Please help me find this species.
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Respected Lane Allen,
Greetings! Thank you for your support and help. I will following your suggestion for future diatoms research.
By,
S. Vijayan
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I want to do liminology of my fish pond. My major concern is with different types of algae present in the pond. ....kindly share your experience regarding sampling of algae and cell count per liter....
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For phytoplankton be use a 20μm net (for qualitiative samples). But for quantitative comparisons you need the volumen of water (the amount of water passing through the collecting net).
In a fish pood if you want quantitative results take samples in bottles, fixed with Lugol’s iodine solution, and you can use the Utermöhl method (sedimentation chambers and inverted microscope).
I hope these information is useful
Maria
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Samples from freshwater, surface water.
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I believe 72 and 287 are Desmids (maybe Closterium closterioides), and 109 is a diatom (Navicula spp.). The filament in the top picture (72) looks like a Spirogira sp.
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Some of the freshwater algae and diatoms we find. But this species identification not easy. Anyone expert help me to find the genus or species?
I am waiting for your support.
Thanking you!
By,
Vijayan
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Maria Van Herk,
Thank you for your help to identifying this species.
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I am analyzing trace metal concentrations in algae pellets, which have been digested in HNO3. Some of the media used to grow the algae have very high concentrations of macro-level nutritional elements i.e. Na, Mg, P, S, and Ca. The range of concentrations is from 1000 ppm to 10,000 ppm. Can I run these samples directly on the ICP-MS without damaging the mass specs/multiplier?
I expect issues like signal suppression and deposits on the cone, but the main concern is not damaging the ICP-MS.
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If the salt load proves too great you can resort to the well-tested, if labour intensive, technique of selective extraction
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In a study in which we want to study the rhodophyte Gracilaria vermiculophylla with PAM fluorometry (diving PAM) we are facing the problem that this small tubular algae doesn't cover the complete area of the dark leaf clip. How can we correct the measurements?
  • Is it correct to add more algal biomass until the area that is measured is approximately covered up uniformly?
  • Should we relativize the output to the area that was surveyed when we take measurements from a single thallus?
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Hi Hector,
In my understanding, the nature of the problem depends on what information you are trying to gain from the PAM. If you are just trying to measure relative parameters (i.e. rETR, NPQ and others) then assuming your background light is constant you should be able to get measurements. As Bjorn pointed out you may have to optimize the F0. The assumption that the background light is constant my be a poor assumption especially if your clip doesn't sit well on the macro algae. I would recommend doing a some tests to see how constant the red background light is. If the background light is not constant I would consider adding aluminium foil or something else around the clip to blockout background light.
If you are trying to measure absolute ETR, measuring of macroalgae absorption will be tricky. Here you may have to find a way of limiting the area of the clip to a know area that you are sure is completely covering the algae biomass.
Hope this helps and best of luck
-Boomie
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Currently, I use pure water as my eluate. I wanted to use NaN3, but our company won't allow me to use that due to safety issues, so I wonder if there is something else I can use to prevent algae growth?
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