Microalgae - Science topic

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.

refer:

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=ε⋅c⋅l

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!

Vasilis Andriopoulos Thank you for your precious answer!

May answer your question.

Best regards

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.

Biologically, I guess that the system needs to be aerobic with as much CO2 as it will otherwise stand.

@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:

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.

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

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.

Please see the attachment

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:

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.

You may try filtration instead: https://lab.plygenind.com/centrifugation-vs-filtration-whats-the-difference-and-how-to-choose-for-your-separation-process

@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:

To prevent or manage excessive green algae growth and mitigate potential risks to the shrimp larvae, you can consider the following measures:

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.

Dear friend Senthilkumaravelan Ayyar

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

https://www.researchgate.net/publication/303976897_Using_Microalgae_for_Treating_Wastewater?

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:

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

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

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).

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].

[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

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

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

Hi 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.

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.

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

for microalgua .you must autoclave all material before using

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.

From what I have read cultivation on an acidic medium totally prevents the infection.

Check:

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.

You will find a similar example in the manual of my software (red intensity of coffee vs concentration):

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.

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

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

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.

According to the shape, it maybe one of the species in Euglenophyceae. Please refer to Eutreptiella sp.

However, I don't like to be a pessimist without ignoring the fact that time presses.

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

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.

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!

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.

ribulose bisphosphate carboxylase oxygenase

Ionically gelled alginate can be dissolved by treatment with chelating agents for divalent cations such as citrate and ethylenediaminetetraacetic acid (EDTA) or hexametaphosphate

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.

Dear Jess,

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

and here:

Best wishes

Eleftherios

Pyro Farms, highly recommended

Joel

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

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

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

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

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

Thank you Gilbert for your valuable feedback regarding this!

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

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

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

Dear Maysan Nashashibi , thanks for sharing this interesting question. It is always a good idea to use the searching box into RG website, often you will get some very valuable informations such as those provided by Aarif 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.

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

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

Hello Alejandra Vásquez

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

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

Best wishes,

Ritesh

As many people have answered, yes you can.

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

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

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

Kindly check also the following useful RG link:

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

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

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

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

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

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

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

hello dear,

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

Hey Maximilien,

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

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

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

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

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

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

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

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

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

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

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

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

Hope this helps, at least for the moment.

Best regards

Niklas Zell

Hello Ruqian,

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

Kind regards,

Ritesh

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

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