Science topic

Microalgae - Science topic

A non-taxonomic term for unicellular microscopic algae which are found in both freshwater and marine environments. Some authors consider DIATOMS; CYANOBACTERIA; HAPTOPHYTA; and DINOFLAGELLATES as part of microalgae, even though they are not algae.
Questions related to Microalgae
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Most of the articles suggested HPLC for the analysis of mycosporins from microalgae, but due to the need for strain screening, I need a rapid method of extraction and measurement by spectrophotometry.
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Dear friend Saeede Taherpanah
Now, let's dive into the world of mycosporin-like amino acids (MAAs) with my fervor!
For a quick method of extracting MAAs from microalgae with a primary measurement using spectrophotometry, you Saeede Taherpanah can consider a simple and rapid acidified extraction method. Here's a brief guide:
### Quick MAA Extraction Method:
**Materials Needed:**
- Microalgae samples
- Methanol
- Acid (e.g., hydrochloric acid)
- Water
- Spectrophotometer
**Procedure:**
1. **Harvest Microalgae:**
- Collect microalgae samples during the exponential growth phase.
2. **Sample Preparation:**
- Wash microalgae with distilled water to remove salts.
- Freeze the samples and lyophilize to remove excess water.
3. **Acidified Methanol Extraction:**
- Add a known quantity of microalgae powder to a vial.
- Add methanol (typically 80-100%) acidified with a small amount of hydrochloric acid (0.1-1%) to the vial.
- Shake vigorously for a few minutes.
4. **Centrifugation:**
- Centrifuge the mixture to separate the cellular debris from the extract.
5. **Spectrophotometric Measurement:**
- Take a small aliquot of the supernatant.
- Measure the absorbance at a wavelength specific to the type of MAA present (typically around 310-360 nm).
6. **Calculation:**
- Use the Beer-Lambert Law to calculate the MAA concentration:
A=εcl
where
- A is the absorbance,
- ε is the molar absorptivity of the specific MAA,
- c is the concentration, and
- l is the path length.
**Notes:**
- Calibration curves with known MAA standards can help in quantification.
- The choice of acid and its concentration can influence the extraction efficiency.
This method provides a rapid way to screen strains for the presence of MAAs using spectrophotometry. Keep in mind that it might not give the same precision as HPLC, but it's a trade-off for speed.
Now, go forth and unravel the mysteries of microalgae with my spirit!
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Hi there. Does anyone know if microalgae (chlorophyta in particular) can use EPS (or polysaccharides) as source of energy? Can you reccomend me any paper where polysaccharides have been used as energy source (with or without success)?
Background of the question -> i'm trying to undersand if algal species in an only-algae consortium may benefit, in terms of energy, from polysaccharides released by other species.
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Vasilis Andriopoulos Thank you for your precious answer!
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Is it positive or negative relationship?
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  1. capsule/ fresh/dry and or...
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It could utlized as feed additives in the form of capsule or just ina minute amount as it has very high profiles of major nutrients and micro-nutrients.
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I am trying to grow microalgae. Between bubbling in air (just air, no concentrated CO2) and shaking, does either of them help grow algae faster? Or do they essentially have any fundamental differences between them?
Also, does anyone have any recommendations for set ups to bubble in air or CO2? (I am in the US)
Thank you!
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Biologically, I guess that the system needs to be aerobic with as much CO2 as it will otherwise stand.
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I got a liquid starter culture of Thalassiosira Weisflogii microalgae. While the starter culture grows indoors. But it does not grow on agar plates using the F2+si medium. I am seeking reasons for its inability to grow on the petri dishes and solutions to this issue.
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@all The inability of Thalassiosira weissflogii microalgae to grow on agar plates with F2+Si medium, despite successful growth in a liquid starter culture indoors, could be due to several factors. Let's explore some possible reasons and potential solutions:
  1. Agar Quality: The quality of agar used in agar plates is crucial. Ensure that you are using high-quality agar and that it's properly prepared. Agar that is not properly dissolved or sterilized can inhibit microalgae growth. Try using commercially available agar specifically designed for microbiological purposes.
  2. Medium Composition: Double-check the composition of your F2+Si medium. Any errors in preparing the medium can hinder growth. Make sure you're following the correct recipe, including the appropriate concentrations of nutrients and trace elements.
  3. Sterilization: Ensure proper sterilization of the agar plates and medium. Autoclave the medium and agar plates at the correct temperature and duration to eliminate any potential contaminants.
  4. Temperature and Lighting: Microalgae are sensitive to temperature and lighting conditions. Confirm that the temperature and lighting in your lab are consistent with the conditions in which the microalgae were successfully grown indoors.
  5. Nutrient Availability: Check if the nutrients in the F2+Si medium are accessible to the microalgae on the agar plates. Agar can sometimes form a barrier that prevents microalgae from accessing nutrients. Consider pouring a thinner layer of agar in the plates to ensure better nutrient diffusion.
  6. pH: Microalgae can be sensitive to pH levels. Ensure that the pH of the agar medium is within the suitable range for Thalassiosira weissflogii. Adjust the pH if necessary.
  7. Inoculation Density: The initial inoculation density can affect growth. Ensure you are transferring an adequate number of microalgae cells from the starter culture to the agar plates. Too few cells may not be sufficient to establish growth.
  8. Contamination: Verify that your agar plates are not contaminated with unwanted microorganisms that may be outcompeting the microalgae. Sterilize all equipment and work in a clean environment.
  9. Adaptation: Sometimes, microorganisms require adaptation to a new growth substrate or environment. Try streaking the microalgae on fresh agar plates periodically to see if they eventually adapt and grow.
  10. Subculture: If the microalgae on the agar plates are not growing, consider subculturing them into fresh liquid medium. Once they are actively growing in liquid culture, you can attempt to transfer them back to agar plates.
  11. Consultation: If the issue persists, consider consulting with colleagues or experts in algal cultivation or microbiology. They may provide specific insights into growing Thalassiosira weissflogii on agar plates.
Remember that microalgae cultivation can be sensitive, and troubleshooting may involve several iterations to pinpoint the exact issue. Careful attention to the factors mentioned above and patience in experimenting with different conditions should help you successfully culture Thalassiosira weissflogii on agar plates with F2+Si medium.
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In D. salina in which beta carotene is accumulated throughout the cells, staining with iodine reagent will cause the whole cell to be stained.
NOTE: These pictures are copyright. No reproduction without written permission from the publisher.
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Here is a paper I recently read:
Salinity impairs photosynthetic capacity and enhances carotenoid-related gene expression and biosynthesis in tomato (Solanum lycopersicum L. cv. Micro-Tom). PeerJ, 8, e9742. https://doi.org/10.7717/peerj.9742
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I want to know about the mechanism through which biological nitrogen fixation occurs in photosynthetic green microalgae apart from previously reported bacterial associations. Like in axenic lab conditions ?
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Typical green microalgae (like Chlorella, Chlamydomonas, etc.) were not known to fix atmospheric nitrogen (N2) directly. They typically assimilate nitrogen from their environment in the form of nitrate (NO3-), nitrite (NO2-), or ammonium (NH4+). The idea of green microalgae fixing nitrogen in the same way cyanobacteria do is intriguing but is not well-established or understood in the literature up to that time.
Cyanobacteria are prokaryotic and are the only group of photosynthetic oxygen-evolving organisms that can fix atmospheric N2.
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I'm looking for recommendations on the best live microalgae to feed Litopeanaeus Vannamei shrimp larvae. Also, if anyone knows the specific Thalassiosira sp. / weisflogii microalgae strain ID that would be suitable, I'd appreciate the information. Thanks in advance for your help!"
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Please see the attachment
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when there's a development of harmful algal blooms which produce toxins such as microcystin, anatoxin, saxitoxin which hamper the ecosystem and due to algaecide treatment these toxins are released in the environment and kill many aquatic organisms I want to check what will happen when both toxins and algaecide are exposed to microalgae. Due to eutrophication there's variation in abiotic factors such as light, temperature, nutrients level, Ph etc I want to know the combined effect of these stressors on microalgae used in wastewater treatment.
I am research scholar interested to work on this aspect. Your suggestions will be very much helpful for me to pursue my research work.
Thank you.
Divyadevale.
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When eukaryotic microalgae are exposed to different abiotic and biotic stresses simultaneously, their physiological and biochemical responses can be complex and varied. These stresses can include environmental factors (abiotic) such as temperature changes, light intensity, pH fluctuations, salinity variations, nutrient limitations, and oxidative stress, as well as biological factors (biotic) such as competition with other microorganisms, allelopathy, and predation.
The specific responses of microalgae will depend on various factors, including the species of microalgae, the intensity and duration of the stresses, and the availability of resources. Here are some possible outcomes and responses when microalgae face combined abiotic and biotic stresses:
  1. Growth Inhibition: Exposure to multiple stresses can lead to growth inhibition or reduced growth rates compared to optimal conditions. This is often due to the diversion of energy and resources towards stress response mechanisms rather than growth and reproduction.
  2. Altered Pigment Composition: Microalgae may adjust their photosynthetic pigment composition in response to changes in light intensity and quality, as well as nutrient availability. These adjustments aim to optimize light absorption and protect the cells from excessive light or oxidative damage.
  3. Cellular Damage and Oxidative Stress: Simultaneous exposure to abiotic and biotic stresses can lead to the generation of reactive oxygen species (ROS) within the cells, resulting in oxidative stress. This can damage cellular components, including lipids, proteins, and DNA.
  4. Enhanced Synthesis of Stress-Related Proteins: Microalgae may upregulate the synthesis of stress-related proteins, such as heat shock proteins and chaperones, to protect cellular structures and maintain protein homeostasis under stressful conditions.
  5. Allelopathic Effects: Some microalgae release chemical compounds with allelopathic effects, inhibiting the growth of neighboring microorganisms. These allelopathic interactions may intensify under stress conditions, impacting the structure of the microalgal community.
  6. Changes in Cell Morphology: Microalgae may undergo changes in cell size, shape, and morphology as an adaptive response to cope with specific stress conditions.
  7. Nutrient Uptake and Utilization: Microalgae may adjust their nutrient uptake and utilization strategies under combined stress conditions to optimize their survival and growth.
  8. Shift in Metabolic Pathways: Under stress, microalgae may alter their metabolic pathways to produce and accumulate specific metabolites that act as osmoprotectants, antioxidants, or other stress-tolerance molecules.
  9. Changes in Cell Signaling: Different stress response pathways in microalgae involve complex cell signaling cascades that may intersect and interact when multiple stresses are present.
It is essential to note that the response of eukaryotic microalgae to combined stresses is highly diverse and specific to each species. Additionally, some microalgae are more resilient and adaptable to multiple stressors, while others may be more sensitive, leading to potential shifts in microalgal community composition and ecological dynamics in aquatic ecosystems. Understanding these responses is crucial for studying the impacts of environmental changes on microalgal populations and ecosystem functioning.
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Dear colleagues and friends
I have encountered a problem with my Microcystis culture. I tried isolating its protein and centrifuging it to collect the biomass. But the biomass won't settle down even after centrifuging at 14,000 rpm for 15 mins. There were always cells floating on the top of the centrifuge tube (I have tried several tube sizes; 15 ml, 5 ml, and 1.5 ml, and several rpm speeds from 9,000 to 14,000). Microcystis may have a gas vesicle so it does not sink. Any ideas about collecting the biomass without breaking the cells?
Thanks
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Hi everyone,
I would like to test the effect of some microalgae (as a source of bioactive compounds) on lipid accumulation in 3T3-L1 adipocytes.
My concern is how handle microalgae and treat cells; due to their characteristics, specially as unicelular organisms, microalgae can be added to the incubation media of cells, and it seems that there are not afected after 24 hour treatment with a high dose. However, if there is microalgae cell wall all the bioavtive compounds remain inside them, is ¡n´t it?
I would be very gratfull if you could recommend me if it is better to make a cell disruption and ad this to the incubation media,
or
If it could be better to try an extraction with solvents.
It´s my first time working with microalgae in cell culture, could you help me? any specific protocol?
Thank you in advance
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The treatment is mention in my research read it how do cell determine its size to grow
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What Causes the green color change? Because Diatoms are produce brown color tint in the tank. Does It lead high mortality rate in shrimp post larvae in larval rearing tank
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@all Apologies for the confusion in my previous response. You are correct that Thalassiosira weissflogii diatoms typically produce a brown color tint in the water rather than a green color. I apologize for the oversight in my previous explanation.
In the context of shrimp larval rearing tanks, the presence of a green color in the water is more commonly associated with the overgrowth of green algae, such as species from the genera Chlorella, Nannochloropsis, or Tetraselmis. These green algae can proliferate under favorable conditions and lead to the water turning green.
Excessive green algae growth can have various impacts on the larval rearing tank and the shrimp post-larvae (PL). While green algae themselves are not usually directly harmful to shrimp larvae, their overgrowth can indirectly affect the larvae and potentially lead to high mortality rates. Here are some possible reasons for the negative effects:
  1. Reduced oxygen levels: Dense algal blooms can deplete oxygen levels in the water, leading to hypoxia or low oxygen conditions. Shrimp larvae require sufficient oxygen for their growth and survival. If oxygen levels become critically low due to the excessive growth of green algae, it can result in stress and mortality of the PLs.
  2. Changes in pH and alkalinity: Algal blooms can alter the pH and alkalinity of the water as they consume carbon dioxide during photosynthesis. Rapid changes in pH can stress the shrimp larvae, affecting their physiological processes and increasing mortality rates.
  3. Competition for nutrients: Green algae compete with the shrimp larvae for nutrients in the water, particularly nitrogen and phosphorus. If the algae outcompete the larvae for these essential nutrients, it can negatively impact the larvae's growth and development.
To prevent or manage excessive green algae growth and mitigate potential risks to the shrimp larvae, you can consider the following measures:
  1. Nutrient control: Monitor and manage nutrient levels in the water, particularly nitrogen and phosphorus, to limit algal growth. Properly balanced nutrient inputs can help prevent excessive algae proliferation.
  2. Light control: Adjust the lighting conditions in the larval rearing tank to prevent excessive algae growth. Algae require light for photosynthesis, so reducing the light intensity or using shorter lighting periods can help control their population.
  3. Filtration and water exchange: Implement an appropriate filtration system to remove excess algae from the water. Regular water exchanges can also help dilute the algal population and maintain water quality.
  4. Monitoring and management: Regularly monitor water quality parameters and observe the behavior and health of the shrimp larvae. If excessive algae growth occurs, take appropriate actions to mitigate its impact, such as adjusting nutrient levels, increasing filtration, or implementing additional water exchanges.
By maintaining optimal water conditions and preventing the overgrowth of green algae, you can help reduce stress on the shrimp post-larvae and minimize potential mortality rates.
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What causes the Shrimp larval rearing tank culture water turns into GREEN COLOR ? which has (live Thalassiosira weissflogii microalgae& industry standard Probiotic & other growth minerals). Also Vorticella infestation problem occurred it leads to high mortality rate in shrimp early post larval stage. Any suggestion to prevent/ reduce VORTICELLA in larval rearing tank
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The green coloration of the shrimp larval rearing tank water can be attributed to the presence of excessive algae growth, particularly the live microalgae Thalassiosira weissflogii in your case. Algae bloom occurs when there is an abundance of nutrients, such as nitrogen and phosphorus, in the water, providing favorable conditions for algal growth. The high nutrient levels can be a result of excess feeding or inadequate water exchange and filtration in the tank.
To address the green water issue, you can consider the following measures:
1. Adjust feeding practices: Ensure that you are providing an appropriate amount of feed to the larvae and avoiding overfeeding. Excess feed can contribute to the nutrient load in the water.
2. Optimize water quality parameters: Monitor and maintain proper water quality parameters, such as temperature, pH, salinity, and dissolved oxygen levels. Regular water exchanges and filtration can help dilute and remove excess nutrients.
3. Enhance water circulation: Improve water circulation and aeration in the tank to disrupt algal growth and promote better water quality. Consider using appropriate pumps or air stones to enhance circulation.
4. Use UV sterilization: Incorporate a UV sterilizer into the filtration system to control algae growth. UV light can help eliminate algae and reduce the green water problem.
Regarding the Vorticella infestation issue, Vorticella is a common ciliate protozoan that can attach to surfaces in the water, including the larvae. It can cause harm and lead to high mortality rates. To prevent or reduce Vorticella infestation, you can consider the following strategies:
1. Maintain clean surfaces: Ensure that tank surfaces, including tank walls and equipment, are properly cleaned and free from debris or organic matter where Vorticella can thrive.
2. Improve water quality: Implement proper water quality management practices, including regular water exchanges, filtration, and maintenance of optimal water parameters, to create an environment less conducive to Vorticella growth.
3. Use appropriate treatments: Consult with aquatic health professionals or experts to identify suitable treatments or additives that can help control Vorticella infestation without harming the shrimp larvae or other beneficial organisms in the tank. These treatments may include specific medications or natural remedies.
It is important to note that specific recommendations and approaches may vary depending on the specific species of shrimp, local conditions, and available resources.
In addition, my published articles can be of interest to you
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What percentage of glycerol is suitable for storing microalgae in -80C for long term purposes?
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dear friend Ayusmita Ray
When storing microalgae in glycerol stocks for long-term purposes, a commonly used glycerol concentration is 10-20% (v/v). This concentration helps to protect the microalgae cells and preserve their viability during freezing and storage at -80°C.
Here are a couple of references that discuss the use of glycerol stocks for microalgae preservation:
  1. Pignolet, O., Jubeau, S., Vaca-Garcia, C., & Michaud, P. (2013). Highly efficient cryopreservation of microalgae using solid cryoprotective media. Bioresource Technology, 149, 432-440. doi: 10.1016/j.biortech.2013.09.057
  2. Day, J. G., Slocombe, S. P., & Stanley, M. S. (2012). Overcoming biological constraints to enable the exploitation of microalgae for biofuels. Bioresource Technology, 109, 245-251. doi: 10.1016/j.biortech.2012.04.088
These references provide insights into the cryopreservation and storage of microalgae using glycerol stocks. They discuss the optimal glycerol concentrations and techniques for maintaining the viability of microalgae during long-term storage at low temperatures.
Additionally, you may want to read some of my publications
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I want to conduct experiments with microalgae biodiesel as a part of my ph.d. please provide the details of microalgae oil supplier details that available in India.
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You can get it from indiamart of you can contact some microalagal oil producing companies like: AlgalR NutraPharms Private Limited , Thanjavur, Tamil Nadu; Green magic. Hope this helps
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Hi,
Which form of RuBisCo is present in various microalgae? What is its content?
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The presence of different forms of RuBisCo in microalgae is influenced by their evolutionary lineage and physiological adaptations, making it important to ask more specific questions about particular microalgal groups. For instance, diatoms, a diverse group of microalgae, possess Form I C/D Rubiscos, which are characterised by small subunits with a short βA-βB loop and a carboxy-terminal extension forming a β-hairpin structure (βE-βF loop).
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i found the content of dry weight of euglena gracilis under mixtrophic is much higher than than under heterotrophic. almost 2-fold. is that right?or something wrong happened?
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Hello Rao Yao
Technically, mixotrophic growth should have higher biomass centration than that heterotrophic growth.
Mixotrophic = autotrophic + heterotrophic
Mixotrophic cultivation of microalgae involves the simultaneous utilization of assimilated CO2 and organic carbon to produce biomass and metabolites via both respiratory and photosynthetic routes [1]. This allows microalgae to utilize both inorganic and organic carbon sources, such as glucose, which can enhance biomass yield [2]. In comparison, heterotrophic cultures grow on organic carbon in the absence of light via combined respiratory and fermentative routes [1]. A study on the optimization of culture media for enhanced microalgae production found that mixotrophic optimized media resulted in maxima of biomass and lipid in comparison to that of other cultivation conditions media [1].
Reference:
[1] Photoautotrophic, mixotrophic, and heterotrophic culture media optimization for enhanced microalgae production - ScienceDirect
[2] Frontiers | Editorial: Mixotrophic, Secondary Heterotrophic, and Parasitic Algae (frontiersin.org)
Thanks
Chandan
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Hello phycologists
I was able to isolate about ten isolates of microalgae (cyanobacteria and eukaryotic) from waters polluted by organic matter, the isolation was carried out on BG11 and BBM medium.
The isolates are stored in monoclonal liquid culture, but not axenic, as there is bacterial and fungal contamination.
-my first question: can i go directly to molecular identification without going through morphological identification? "I do not have the expertise for microscopic identification"
-my second question: concerning the molecular identification of microalgae, what are the best primers I can use to identify eukaryotic microalgae and cyanobacteria (could you suggest me some articles).
-My third question: can I run the PCR without having axenic cultures of my isolates, in other words, does the presence of DNA from bacterial and fungal contaminants not influence the PCR results?
Thank you in advance for your answers.
phycological greetings
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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.
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how the nutrient uptake by microalgae grown in wastewater can be measured? method of nutrient uptake in algal biomass?
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Hi Safina,
Microalgae can be taken nourishment in domestic wastewater due to containing the nitrate, phosphate. Other alternative, you can cultivate by chemicals, a host of vitamin. Also, light, CO2 concentration and oxygen to be required for their live.
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I am trying to cultivate fresh water microalgae in BG11 medium but I have difficulties to grow at light intensity 150, it can grow only at 45 μE m−2 s−1.
So, anyone have any idea about I should to do to let it grow at high light intensity?
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Thanks for your answer.
I will try.
But another thing I tried also as the incubator that I am growing in does not contain carbon dioxide and the light intensity is 45 but I split the culture into two flasks then I put one in the infors with CO2 2% and 100 micromole light and the other in CO2 0.2% and 100 also but I found both of them died, from this I thought the reason is light.
but I will try to cover one of them with foil to see if the reason is light or not.
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Hello fellow microalgal researchers. We are looking to set-up a turbidostat system in our laboratory and we are wondering if anyone has recommendations of systems that they are working with?
We have a multicultuvator MC-1000 (PSI) system in our laboratory and are thinking of buying the Turbidostat Module TS 1100 (PSI) that will allow the multicultivator system to maintain turbidostat growth. Does anyone have experience with this system and can comment on its durability, reliability and worth relative to price?
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Look at the turbidostat from Algenuity. Their data collection modules are much better and and bioreactaors are much better designed. British company saw it in US and France conferences. https://www.algenuity.com/algem
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To obtain pure microalgae cultures I have inoculated the water sample on BG-11 and BBM media using spread plate method. I am getting yellow colored colonies on both the media plates. Kindly suggest me some points to avoid contamination in plates.
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for microalgua .you must autoclave all material before using
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In order to measure the oxygen evolution of microalgae cells I have been using an Oxytherm chamber. In order to make results comparable, the same cell desnity of cells had been used. As the culture grows, I need to dilute the samples more and more to which I used MilliQ. Is there a risk that the cells could burst at a 50% dilution?
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Diluting microalgal samples can have an impact on cell viability and oxygen evolution, as it can change the osmotic pressure within the cells. There is a risk that cells could burst at a 50% dilution if the osmotic pressure difference between the cells and the diluent (in this case, MilliQ) is too great.
In general, the osmotic pressure of cells is influenced by the concentration of salts and other solutes within the cells, and the concentration of these solutes in the diluent. If the concentration of solutes in the diluent is much lower than in the cells, water will flow into the cells, causing an increase in volume and potentially leading to cell lysis or bursting.
To minimize the risk of cell lysis, it is important to carefully control the concentration of salts and other solutes in the diluent, and to gradually dilute the cells over several steps to allow the cells to adjust to the changing osmotic pressure. You may also consider using other methods to adjust the osmotic pressure, such as adjusting the pH or adding compatible solutes to the diluent.
It's also important to keep in mind that cell lysis can also be influenced by other factors, such as the age of the culture, the presence of contaminants, and the type of cells being used. To ensure accurate and reliable results, it's important to carefully monitor the cells during dilution and oxygen evolution assays, and to take appropriate measures to minimize the risk of cell lysis.
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Good day All
I am busy cleaning up the microalgae strain Haematococcus pluvialis isolated from my bird bath outside. I have been battling to get rid of the Paraphysoderma sedebokerense (fungal parasite) on this strain for a very long time. Is there perhaps someone in this community with more experience who can advise me on which specific antibiotics and / or fungicides would be most effective for cleaning H. pluvialis? Your help will be much appreciated.
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From what I have read cultivation on an acidic medium totally prevents the infection.
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I am using two microalgae strains (Haematococcus pluvialis and Chlorella zofingiensis) which are well known for production of astaxanthin. but after 15-20 days growth of microalgae in TAP medium the coloration of medium is changed to yellow and after analysis in the HPLC the lutein concentration is high as compared to astaxanthin. Is there any possible reason behind synthesis of lutein?
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It's possible that this is due to incomplete carotenogenesis. In reality, the major pigments in green algal cells are chlorophyll a, chlorophyll b, and lutein. When carotenogenesis begins, chlorophyll and lutein breakdown begins. It is possible that you should focus on stressor optimization in the second phase of the cultivation procedure.
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I want to make a correlation between the optical density and dry weight of Chlorella Vulgaris. I have dried specific different diluted volumes of the culture but don't know how to make this correlation and what will be the unit used in this correlation.
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You will find a similar example in the manual of my software (red intensity of coffee vs concentration):
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the sample was isolated from polluted river water and cultured in f/2 media according to the European standard. The images are a little bit confusing, so can you help me to id them?
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The images have too little magnification to be able to say anything for sure. With approx. 400X magnification, diatoms look like this, for example: Unidentified alga from fresh water.
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Dear diatomists,
Can someone could say to which Thalassiosira species this diatom belongs? Or it is not a Thalassiosira?
The scale bar is 10 mkm.
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Anton:
It looks like Actinocyclus although I don't see characters like the pseudonodule and others. More details would be needed to properly determine the taxon. Consider Hemidiscaceae in Hasle & Syvertsen in Tomas (1996) and Round et al. (1990).
All the best.
Eugenia
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I have been engineering E.coli for few years, but this is my first time transforming microalgae.
I am trying to transform C.vulgaris, using agrobacteirum-mediated trasnformation method. I found a research that used Freshwater Culture Medium (FCM) containing Bold’s Basal Medium (BBM) major salts and supplemented with F medium-trace metals and vitamins for agrobacterium-mediated transfromation of C. vulgaris (https://doi.org/10.1007/s11274-011-0991-0), but I only have BG11 broth in my lab.
Would using BG11 broth significantly affect the transformation efficiency or would it have nothing to do with culture medium?
Thank you.
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Hi, I have some experience about microalgae culture, If you use BG11 or BBM medium in both cases probably you obtain a good culture, however I suggest you that use another economic alternatives , best regards from Colombia
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Recently, we identified our microalgae using 18s rRNA, and the data showed that the isolate was 99% similar to unclassified microalgae with general information on NCBI and no publications.
Our publication is interested in synthesizing NPs using microalgae. Does it necessary to undergo a classification process, or it is enough to publish the isolate and clearly state that these algae are not classified?
Thank you
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Most of the algal species are described based on morphology. So one would expect the description of its morphology and morphological identification. If you already classified the organism as an alga you should know at least to which class it belongs and how good molecular references are for this class.
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Flagellate located in estuarine waters of Ecuador
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According to the shape, it maybe one of the species in Euglenophyceae. Please refer to Eutreptiella sp.
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Greenhouses gases like carbon dioxide, methane,nitrousoxide,CFC,ozone etc. may be reduced by use of hydroelectric,wind energy,biofuel,solar energy ,microalgae etc. to maintain earth temperature not above 1.5 to 2 degree centigrade from the present mean earth tempearture according to IPCC.
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However, I don't like to be a pessimist without ignoring the fact that time presses.
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I found a published photo where author writes that this is diatom species Hyalosira delicatula. But I have some doubts about such conclusion.
Unfortunatedly I do not have any photographs and the description of Hyalosira delicaula. So, can anyone familiar with the genus Hyalosira confirm or refute this conclusion?
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Dear Anton,
The genus Hyalosira was amended and lectotyped by Lobban et al. (2021); Diatom Genus Hyalosira (Rhabdonematales emend.) and resolution of its polyphyly in Grammatophoraceae and Rhabdonemataceae with a new Genus, Placosira, and five new Hyalosira species, Protist 172, 25816.
H. obtusangula, which is one of the species together with H. delicatula described by Kützing (1844), was chosen as the lectotype of Hyalosira.
The specimens that you are showing in the pictures clearly do not match Hyalosira emend Lobban, Ashworth and Majewska in terms of girdle band morphology: copulae bearing two rows of areolae separated by a midrib.
Even when certain characters are not in view, the bands bring the displayed material closer to Placosira as mentioned by Roksana in her answer.
All the best.
Eugenia
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Usually, the morphology of Spirulina is a screw-like coil. However, my strain changed its morphology into a straight form. How does it happen? Can it change back into a screw-like coil shape? Any suggestions or advice?
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It is not yet clear why some spirulina cultures are more "linear" than others. Growth conditions are certainly the main cause of the morphological change of spirulina, but sometimes it is also due to a specific growth phase of the biomass.
I would check if there are any problem with culture media, or if is present a contamination by other cyanobacteria.
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I collected various samples from the coastal region, which will be the best media to culture the microalgae from that sample under a laboratory scale.
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Dear Sujit,
Patiently, is probably the best way! Unless you have a cell sorter. You will need to continuously carry out single cell isolations in a well plate moving the cell of interest into clean, appropriate media under species specific optimal conditions until that cell divides to become the dominant strain. At that point you may be able to carry over more than one cell into a fresh well. Then, it may be necessary to treat the new culture with antibiotic or anifungal agents. Sadly, it will not always be possible, as many species will not tolerate the conditions of single cell culture. Best of luck with it!
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I had grown Chlorella sorokiniana in BG-11 media supplementing with antibiotic and antifungal cocktail. The culture became healthy green in a few hours to pale yellow in a day. Suddenly, in next day, the culture converted its colour from pale yellow to white with some floc-formation and after a day or two colour changed to green again . So what should be the reason of sudden colour changes in the micro-algae culture?
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Sheetal Parakh hello Sheetal Parakh, Actually my hypothesis behind this is its acclimatization patterns towards nutrients present in the media, or I can say it took its own adaptation time or pattern. Maybe you can experiment more with changing the ratio of nutrient doses especially N: P. I hope this will help.
Thanks.
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Hello friends. I want to extract rubisco from spirulina microalgae does anyone have a solution? Do you have a proper protocol?
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ribulose bisphosphate carboxylase oxygenase
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Hello everyone,
I am going to prepare either an alginate or carrageenan hydrogels for immobilization of microalgae. The microalgae will be expected to be retrieved from the hydrogels after finishing cultivation. In that case, hydrogels need to be destabilized/dissolved. Can anyone suggest any chemicals or processes for this ? I found some information about chelating agent like sodium citrate or EDTA as chelating agent for Ca2+ ion in alginate matrix, but not sure any other agents out there are available, especially for K+ ion.
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Ionically gelled alginate can be dissolved by treatment with chelating agents for divalent cations such as citrate and ethylenediaminetetraacetic acid (EDTA) or hexametaphosphate
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For some reason, I had no choice but to culture my microalgae in the same LAF that was being used for fungal studies as well. Now for obvious reasons I have started getting fungal contamination in my microalgal plates. I have changed the LAF and even on subsequent re-streaking, I am unable to get rid of it. Any suggestions as to how to make the microalgae free from fungal contamination?
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Hi,
Common growth media for microalgae generally do not support heterotrophic microbes as they lack assimilable carbon sources, and the same is true for Bold media and its 3N version. It appears that natural cell lysis with growth is enriching your algal cultures with nutrients required to support contamination. Lysis is always almost negligible, but because you are using a common culturing facility, it is making your cultures more susceptible to contamination.
As algae are very slow-growing, even minute contamination is supposed to outgrow soon. Thus it is important to analyze your system and practices in more depth to suggest some measures and tricks to avoid any contamination.
All the best for now.
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This algae invade my culture lately (Chlamydomonas). It will caused the culture to clump and eventually die. This contaminant was motile, with more than 4 flagella observed.
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Dear Jess,
probably it is Poterioochromonas, a golden algae. You can find some information here:
Best wishes
Eleftherios
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Hello...
Two months ago, I bought pyrocystis fusiformis culture for my research. Unfortunately, no bioluminescence was observed, and the culture seemed to be deficient from the start.
Now I'm trying to buy a Pyrocystis fusiformis culture again and found a lab called PyroFarms, but before I order, I wanted to check if anyone had any past experience with them. or alternatively, if you could name any other trustworthy sources where I could buy a P. fusiformis culture, I would be grateful.
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Pyro Farms, highly recommended
Joel
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Hello everyone. Hope this question find you well.
I was recently reading about fatty acids because my master thesis is going to be about them and the most of their perspectives, like a review. And a question arose since I read in a book that there are some trans-unsaturated fatty acids found in nature (microalgae, bacteria, plant, etc), I can assume why and their reasons to be, but my question is that if these trans-unsaturated fatty acids can turn in cis fatty acids because some enviromental condition, as a reduction in the environmental stress to which it may have been subjected the sample.
I was searching for some information that can answer my question, but nothing at the moment. I'll appreciate so much your help with this. Thank you!
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Yuvraj Saxena thank you for your answer! it truly helped and brought me a little more information about it
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I found this specie in estuarine waters of Ecuador
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Of course not! Strictly speaking, it is a nomenclatural and spelling error. But in the case of Scenedesmus, Chlorella, Chlamydomonas, Oedogonium, and some genera that begin with compound Latin letters (Oe, Sc, Sch, etc.), this spelling was used to avoid confusion with other genera in the floristic lists. Sorry, I should have written S. indicus and S. producto-capitatus.
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Good afternoon, I am working with the production of Chlorella sp. in industrial waste and I would like to know what parameters are the most optimal to evaluate in this industrial waste... If anyone has any information about it, I would appreciate your contribution.
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pH, NPK, C content, DO, moisture. If you need any more information then write back to me
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This one is a marine microalgae isolated from Bay of Bengal.
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Can you provide more clear views
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This is Haochen Zhang, a MA design products student from Royal College of Art. And I’m currently working on a project about removing nutrition of flashed dairy manure. After learning some relevant materials, I realized that microalgae is a very effective means of treating wastewater, and I want to design products which can take use of microalgae (Chlorella) to help local rural residents transform the water to new resource. But now I lack of detailed and professional guidance. So I sincerely hope that I can get an opportunity to learn from you, and get more advice about this project.
Here are some questions which have been confusing me so much:
1. Will sterilization influence the effect of chlorella growing? If I want to sterilize the flushed manure water (100% sterilization), how much do I need to heat it to what temperature? And how long should it last? Can a sterile environment be achieved by simple boiling? Will high-temperature heating produce toxic substances in manure?
2. Is it necessary to re-dilution to feed chlorella with flashed manure water? Is it sufficient as a source of nutrition for chlorella? Do I need to add more nutrients? To what extent can the nitrogen and phosphorus in the water body be absorbed by chlorella?
3. What indicators should be paid attention to in the environment of cultivating chlorella? For non-professionals, is it difficult to cultivate chlorella with the water? What is the most suitable container volume for cultivating it?
4. Can the cultivated chlorella be directly used for fertilization (spraying or watering on vegetables)? Does it have a positive effect or additional economic value for agricultural production?
I sincerely hope that I can harvest precious opinions and opportunities to learn from you. Your suggestions must be very valuable to me. I hope with your help, my project can become a good design that has a practical effect on society! If possible, I hope to have more contact with you, thank you very much! !
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Arne Andersen Thanks for your suggestion, it seems to be a more efficient and practical way. One thing I'm not sure about though is whether it takes longer to degrade cow dung water by the growth of saprophytes than when using microalgae?
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Despite the high quality of fish-based ingredients in aquafeeds, reported unsustainability caused by their use in aquaculture, have raised global concerns and effort for replacing them.
Among the potential candidates ( insect meal & oil vs plant meal & oil vs microalgae meal & oil : Spirulina , Chlorella, etc. ), which ones have no side effects or don’t generate new environmental impacts ?
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Microalgae considered a a noble biomass for production of high value products, food and feed, Pharmaceuticals as well as Biofuels.
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Thank you Gilbert for your valuable feedback regarding this!
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How does the F/2 medium help the growth of the Dunallila salina microalgae?
why do we use this medium?
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This medium contains abundant levels of N and P (also Si for diatoms and other Si-requiring species) along with a comprehensive set of trace metals and vitamins. The N:P:Si ratio is close to the classical Redfield ratio. As such, the medium is very effective in supporting the growth of most algae. Another similar medium is L1, which has similar recipes with small modifications of trace metal composition.
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What is the main mechanism of wastewater treatment through microalgae in open ponds and closed cultivation? How does it work when there is no oxygen and carbon dioxide in closed cultivation?
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I agree with Vit, in open ponds, there is CO2 whereas in closed at the beginning they will use organic carbon, but if there is light they produce oxygen and co2 in the dark phase o photosynthesis, so you will have mixed metabolism. Microalgae are supposed to remove nitrogen and phosphorous from wastewater, but they can also remove carbon depending on growth conditions.
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I'd like your advice on Pyrocystis Fusiformis growth. I purchased a culture a month ago and began cultivating it right away. Things, on the other hand, do not appear to be moving forward. There has been no bioluminescence observed yet, and the algae's growth has been poor. The culture was maintained at a pH of 8.2 and a temperature of 20 degrees Celsius with a 12-12 dark/light cycle. Cell counting under a microscope was used to monitor the algae's progress, but because I don't have much experience in the field of microbiology I wasn't sure of my count. I've attached a photo of what I see under the microscope, and I'm not sure if it's algae cells or not.
I'm hoping you could give me some suggestions on how to improve the growth of algae and its bioluminescence.
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Barry C Smith F/2 media was utilized, along with a quite bright LED light (5000 lux measured using a lux meter app on the inoculum surface). The culture volume is 4L, while the inoculum volume is around 6L.
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Hey all !
I am looking for the best method to prepare microalgal slides for TEM characterization, especially as I am working with nanoparticles.
Thanks all in advance :)
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Dear Maysan Nashashibi , thanks for sharing this interesting question. It is always a good idea to use the searching box into RG website, often you will get some very valuable informations such as those provided by Aarif Hussain Shah . Specially in the first link you can see a good TEM sample preparation protocol for bio-samples with nanoparticles.
That is a generic protocol, and it must be adapted to each sample and to what you want to characterize. For example, it will be different if these nanoparticles (NPs) are in the liquid media, on the microalgae´s surface, or dispersed into the microalgae´s tissues. Problems associated to NPs aggregation will probably happen in the two first scenarios, when the NPs are dispersed into the liquid media and by evaporation of it, they end up on the surface of the algae. Some NP´s could be attached to the algae surface and evaporation of the media could add more from the media. If you are interested about the NP´s already bonded to the microalgae surface, and there are NP´s dispersed into the media around, it would be useful to remove the algae from the media, wash them up with some clean media (bear in mind that this clean media should not remove the bonded NPs from your algae), then dry the algae as explained in the previous links or if it is available, use a TEM cell for liquid samples. Of course, the microalgae must be thin enough to work in transmission mode as pointed by Mohammed Amer Shaheed .
If the microalgae have the NPs inside their tissues, the chances of aggregation by drying are very few, the algae tissues will keep them apart, so that if you finally observe aggregation of NPs, it means -generally- that they were already aggregated before the drying process. Once more, Mohammed advice is key, you will need a very thin section of your algae to let the electrons go through the sample and forming a image. You can get such thin sections embedding the sample into a resin and cutting it with an ultramicrotome. Otherwise, you could use a FIB (Focused Ion Beam), a very focused and thin beam of ions to cut a lamela (a very thin section) of your sample, which is then placed on a TEM holder grid with the help of micromanipulators.
TEM works by transmission, transmission of electrons through the sample, so depending on the density of each part of your sample (the value of Z, the atomic number of the atoms on each part) you will see more or less contrast. For example, if your NPs are metallic, they will be denser than the organic tissue of your microalgae, and a good contrast will be expected. But if your NP´s are light, say carbon NP´s or Si NPs the contrast with the organic matter will be smaller.
It could be very useful, before to go to the TEM, to observe the samples with an optical microscope (may be a confocal, looking for some fluorescence from the NP´s), or a faster option, to use a SEM, specially if you are studying NP´s on the algae surface. Normally you would need to dry up your sample as with TEM, but here -with SEM- you don´t need to worry about sample´s thickness too much. SEM often requires conductive samples or coating your sample with a very thin conductive layer (gold, carbon, iridium...), but modern equipment also let us to use the sample uncoated, thanks to charge removal methods.
Hope this helps. Good luck with your research work and my best wishes.
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I want to know the pro and cons of this production?
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Micro algae is a better option. However, you have to cultivate the correct species. Otherwise oil yield will be low.
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Good morning, I have a question and I hope someone can help me. I have as inoculum chlorella vulgaris, with photoperiod of 12/12 hours, shaker agitation up to a volume of 500 mL and agitation provided by fish tank pump for volumes of 1 L and up.
I have observed the formation of a foam on the top of my crops, which I associate to a bacteria producing it but I have not been able to control that, any recommendations?
Also in the upper part of my containers it is forming like a green slime where my microalgae are staying and although I try to homogenize them they do not unite. Could this be associated with
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It may due to the saponins produced by Chlorella. Generally they exhibits high phytochemical content especially n log phase. please subculture the same and if the property reappear go for saponin test
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Good morning, someone could help me to decipher what is causing this in my solid media, I planted chlorella vulgaris in plate count medium and in nutrient medium and I observed that some of them became fuchsia with the passing of lso days, could it be some metabolite or some microorganism?
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Maybe your plates are contaminated because changes in pH or growth of the microorganism may lead to a change in colour. Also, may I ask what agar are you using and exactly how long have these plates been inoculated. If your cultures were green at first and then turned pink, then it just means that the culture had become old.
To be sure, I suggest you cut a piece of the agar and grind it in some distilled water. Then observe the contents through a microscope. You might get an idea of what happening in case its a contaminant.
Best wishes,
Ritesh
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Can I extract carotenoid pigment from microalgae (C. sorokiniana and D. salina) without freeze-drying?
I already searched and read references about drying process that we can use heat-drying method but that wasn't recommended because could cause degradation of carotenoid content. So if I don't have a freeze-dryer, can still extract the pigments from wet biomass?
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As many people have answered, yes you can.
If you want to see the amount of carotenoids per weight, you can determine the dry weight per culture medium, and separately determine the amount of carotenoids in the wet biomass per culture medium.
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Hello,
For a project aiming to produce Spirulina production for human use, I am lokking for a pilot scale (around 10 or 20 thousand liter capacity) PBR supplier..
I will appriciate if you suggest a good company producing LED light integrated tubular PBR in Europe or Asia.
thanks alot for your help
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Thank you for all of yours suggestions, Schott proveide really high tech products but it is really expensive. BBI is no longer produce reactors in pilot or industrial scale. I didnt know algoliner, I will contact them. Thnaks again.
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I recently published an article "Shining a Light on Wastewater Treatment with Microalgae" doi.org/10.1007/s13369-021-06444-3 https//rdcu.be/cE0C2 and now I would like to prepare a much shorter version of the article for publication in Frontiers for Young Minds. There are no publication charges, and this is probably the best way of communicating with the next generation of scientists. Because the article is intended for younger students the journal limits the length of manuscripts to 1500 words, 3 figures, and 5 references. The scope of the article will be determined by the team that agrees to collaborate on this venture. It is not required that authors are well versed in wastewater treatment, microalgae, or renewable energy. Rather it is only required that you have an interest in these topics and are willing to collaborate.
Can you help?
John Kilbane
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Jacob de Feijter, thanks for your interest in collaborating on this topic. The task has already begun and correspondence among collaborators is taking place via email. You may contact me at kilbane@iit.edu and we welcome your involvement.
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We are starting soon an in vivo trial with broilers fed diets supplemented with microalgae and insect full fat larave. We will collect blood samples and I'd like to have your suggestions about the best parameters to be analysed on blood to evaluate the oxidative and metabolic status
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Kindly check also the following useful RG link:
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I m working on biosafety evaluation of GMO microalgae, can anyone guide me which is better and acceptable method (lyophilisation or simple heat drying) for drying as i am going to use it as feed for fish? Also after harvesting pellet of algae, at what temperature i can store before drying, 4 C or -20 C, and for how long without affecting cell integrity?
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I recommend freeze-drying. It preserves all the vitamins and other heat-sensitive micro and macro nutrients of the algae if you use them for fish as feed. If you use freeze-drying, you can store algae in -20 just after harvesting.
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My name is Stanley please help me with the name of the microscope that I can use to view the microalgae structure and able to isolate a single strain.
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I would recommend an inverted microscope with sedimentation chambers, so you won’t have to squeeze your algae between slide and cover glass. This also means you can use magnifications of 630 times, maybe even 1000 with immersion oil. Use both live samples and lugol preserved ones if your specimens are too mobile.
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Actually for my thesis bg11 medium sample in scientific shop are unavailable. So for that any alternative to bg11 medium
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You could prepare BG11 from salts in your lab (www.uni-goettingen.de/de/186449.html)
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Photoautotrophic regime is known to be not as productive as photoheterotrophic regime, mainly due to the fact that in photoheterotrophy microalgae has both light and organic carbon availability. Is there a situation where the opposite happens? What could be the possible explanations? I appreciate your thoughts on this.
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Thank you, Maryam Fath ! Do you have some references on this? Could you please share the link to the papers with me? I appreciate it!
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Hello! I am trying to extract chlorophylls from microalgae, and I would love to ask how can I separate chlorophyll a from chlorophyll b? I want to get each of them separately. Thank you in advance.
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I am running a drying model and would like to have the water content of a chlorella vulgaris microalgae, before drying. Thanks. best.
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Hello. From my experience in the lab, centrifuged microalgae sludge from a 3L Erlenmeyer flask could accumulate a fresh weight of about 2-3 grams and when fully dried would give a weight of about 0.6 - 1.0~ grams. This is a rough estimate as values can vary based on treatment conditions as well. Hope this helps with your project
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Currently I and my group members was tasked on a project on using carbon dioxide as main feed for microalgae product formation. So, we decided on using Chlorella vulgaris but now we are all stuck on balancing the equation since
1. we failed to properly formulate chlorella vulgaris
2. the pathway we have chosen does not show its by products
Without those 2 we cant do economic potential analysis to see whether its profitable for production. any ideas how we should proceed? we may need to change the microbes we use thus the product and that put us back into square one. btw we are trying to get beta carotene as our main product. and the pathway is chosen from. or have we gone the wrong steps?
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You can also try "stress imposed by environmental conditions, such as , temperature, heavy metal concentration, and nitrogen availability; and also UV light stimulation.
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I am making kosaric media using artificial seawater in various salinity for microalgae cultivation. At first, I tried to filter the seawater and then add chemicals. After autoclaving, there was some deposits at the bottom of the media. I also tried to autoclave the artificial seawater and media separately, and then let both solutions cooled down and added together in laminar flow. However, once I added the kosaric media into seawater, precipitation was observed and large amount of white crystalline substance at the bottom. Is there any other way to make it?
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hello dear,
i think you can first filter your seawater, then add your desired chemicals might be these chemicals react with each other i am not sure but i heard from my friend, he face such this type of problem.
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I want to quantify total carbohydrates in Hemerick media ( a media for microalgae growth). After I added phenol & sulfuric acid the sample becomes totally black. This problem isn't observed with artificial sea water media and the same phenol/sulfuric acid solutions.
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Hey Maximilien,
If You have a look at the components, the problem arises from nitrate. Nitrate reacts with sulphuric acid as NO3- -> HNO3 -> NO2+ + H2O.
NO2+ reacts with phenol to nitrophenol, which absorbs in the UV-region and looks dark-greenish.
I was facing the same problem and I tried to solve it as follows (please consider citing me as I cannot find any comparable method):
I measured aqueous nitrate standards and found that they have a plateau around 490 to 510 nm, which is bad as it increases the absorbance exactly at the point, You want to measure the carbohydrates. When nitrate is consumed during the culture, there is a time-dependent absorbance decrease, which renders the results useless.
But there is good news: a plateau means that the slope (1st derivative) in this region is 0, so if you take the smoothed 1st derivative of the spectrum and read the value at 503 nm instead of 490 nm (steepest slope for glucose standards for me), You will get the carbohydrate value You want without the influence of nitrate (since its slope adds 0 to the steepest slope of glucose).
Of course, this implies that You do not use the calibration A(c) = a * c(Gluc) + b as for standard Lambert-Beer-Bouguer, but you need to calibrate dA/d\lambda = a * c(Gluc) + b, which is in perfect accordance with the Lambert-Beer-Bouguer law.
I verified this method in our lab: Nitrate has no influence on the kinetics of the reaction and when I calibrate with aqueous (non medium) glucose standards and measure a glucose standard in Zarrouk medium (similar to Yours), I get a recovery rate of > 95% to 100%.
So, I can assume, this Derivative-Spectroscopy-Method is adequate, but you should post a picture of your spectra (nitrate/glucose in water & glucose in medium, full spectral range) because the assay is very sensitive when it comes to reagent ratios. With this method I fixed the nitrate problem for us, but there is another one. In supernatants from real algae cultivations I get peaks between 500 and 600 nm and since they are not a plateau, they lift the slope of glucose at 503 nm, which dramatically affects my results (lowers them).
I tried to overcome this by taking the smoothed second order derivative at 490 nm (second order derivative is minimum at the top of peak), but since I do not know what causes the colour between 500 and 600 nm I cannot exclude the influence of the colour by another hidden peak below the glucose peak. The influence of nitrate is also 0 for the 2nd derivative since the second order derivative of a plateau is fortunately 0 as well. To sum it up, the new calibration at 490 nm would become d²A / (d\lambda)² = a * c(Gluc) + b (also Lambert-Beer-Bouguer) - but I cannot be certain that this is working in real samples which cause an absorbance at 500 to 600 nm after the reaction with phenol and sulphuric acid (only with both, not with one of them alone).
I've not found anything mentioning the influence of nitrate in the assay except for this (http://www.bioline.org.br/pdf?ja06013), which is soley about nitrate.
I'm really in a need for help what causes the absorbance between 500 and 600 nm, since I've tried everything (various salts, protein (BSA standard), oxidation with persulphate, acidic pre-hydrolysis, cooking, performing the assay with algae biomass still in the sample), but I was completely unable to mimic the additional peaks. I'm growing desparate and depressive about this because I cannot find any literature regarding this. The assay was just not intended to be used in saline solutions and on top of that, nitrate is impossible to remove except for dialysis (cost, effort, etc...).
Maybe we (and I hope experts) can collaborate on this problem. If someone wants to contact me, just google my name + "OTH AW". I have Excel- and Python-software for smoothing, derivation if somebody is not into programming.
Hope this helps, at least for the moment.
Best regards
Niklas Zell
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Hi everyone. I am measuring carbohydrate content of some microalgae. While I was making the standard curve, I noticed that the cultural media (F2 medium and ESAW medium) can also react to phenol and sulfuric acid, leading to a strong yellow color which affects the final result. Do you have any idea why this happens? And how can I solve the problem? Thanks a lot!
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Hello Ruqian,
Saeed and Anthony are correct. It is best to harvest the cells first, wash the pellet and re-suspend in for e.g distilled water. However, I believe it would be a better approach to dry your sample first and then use the dry biomass for the test. It should give more accurate results.
Kind regards,
Ritesh
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I know that this depends on the rheological factors of the medium, but if you know any, please let me know.
Best Regards,
Roberto Novais
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You have to calibrate according to your growth conditions.​ Make​ a​ standard​ caurve and​ compared batween OD and​ cell​ counting
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I am conducting an experiment to obtain enhanced antioxidants production in microalgae by implementing a 3x3 full factorials (three factors each at three levels) abiotic stressors. I would like to analyse my data using minitab software or jmp and obtain optimal levels of factor for maximum antioxidant production.
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You would first need to do a screening study with your 3 factors in jmp, I would suggest a 2X3 (8 runs) with centre points to account for curvature rather than a 3X3 (27 runs). Use the screening study to remove factors that are not significant. Check for the validity of your model by use of graphical methods (residuals and a normal plot) and numerical methods (ANOVA and lack of fit). Once the model is shown to be sound, fit in a linear model based on your significant factors - this linear model can be used to do a method of steepest ascent if your initial screening didn't identify the region of maximum yield - After this, augment your initial screening study with axial points and carry out a response surface study using a central composite design (CCD) or a Box-Behnken design (depends on your factor ranges). The response surface method (RSM) will give you your optimum values by analyzing the prediction profilers, setting desirability to maximize yield and reading response surface curves or contour profilers. Finally, fit in the second-order model.
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I am a PhD student, working with microalgae,
i have a doubt, can microalgae utilise water soluble cellulose like CMC as carbon source?
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