With dilution in a steril petri disc.

If you want to remove the protozoans you will neede to sieve the seawater after 10 um, what you can you need to identify which species you cultuing, isolate them in another petri dish, and if you see other protozoan are still in it, isolate them in another pteri dish, in this way you will have very minimum number of protozoans in the samples...

Best wishes

sonia

Hi Satya,

Here are some basic techniques that are followed to get pure local culture after series of process. Sterile culture of diatom can be obtained by isolation of endemic strains is not simple because of small cell size and association with other epiphytic species.

1. Through serial dilution culture method also it is possible to obtain after 3-4 cultures. 2. Successive plating on agar media and separation using capillary pipettes.

3. In common bacteria and protozoans can be eliminated from phytoplankton culture by washing or plating in the presence of antibiotics. The sterility of the culture can be checked with a test tube containing sea water with 1 g/liter of bactopeptone. After sterilization, a drop of the culture obtained from serial dilution or agar plate culture can be tested by adding to prepared solution and if any residual bacteria is available in culture will turn the bactopeptone solution turbid.

Hope so this will be helpful to find solution for your problem...

Hello,

If possible work in a flow laminae hood under a dissecting scope and pick single diatoms then rinske them in 0.2 micron filtered autoclaved seawater a few times. Heterotrophic bacteria can be kept under control, if a problem, by using a mixture of different antibiotics (that you need to test for mixture composition and concentration). It is rather an intense process, but it is not impossible. Cheers fm

Hi,

Although somewhat comprehensive, you may try to directly isolate your diatom of interest from the field sample using a capillary made from a glass disposable pipet.

Identify the species of interest, suck only this cell in the capillary and transfer it at least two times to a drop of sterile sea water medium. Use a clean capillary everytime. Finally transfer the cell to a tube with sterile medium. Check whether growth occurs.

Since you select one cell, you should end up with a clonal strain. If bacteria occur you can use several antibiotics to get rid of them.

You just need a binocular, pipets, tubes and culture medium.

Hi,

I usually use a combination of liquid and solid media to isolate n strains. By culturing the strain in a rich liquid media with plenty of inorganic carbon you enrich for phototrophs, then plate out some of the culture on an agar plate and selecting isolated colonies. Going back and forth a few times between the plate and liquid usually gives rise to axenic cultures. You can also move colonies or cells on the agar plate to isolate them manually.

You can try with Glass Pasteur pipette. Just take some microlitre of natural water sample in a slide under a microscope and pick your desired diatom species with Pasteur pipette. To avoid contamination just autoclave the pipette leaving that rubber or latex bulb. Here from you can start to do the serial sub-culturing to avoid the contamination by other diatoms. If you are able to pick only some low number of diatom (lets say only 15-20), first place them in small volume of f/2 media and gradually you can increase the media volume also. This is one of the easiest process to establish a pure culture that we do in our lab on a regular basis. Later you can avoid the bacterial contamination (if necessary) by adding some antibiotic. Hope it will help you. Best of Luck!!

Larger planktonic species can be isolated by hand under low power light microscope, using pulled pasteur pipets or even regular micropipet tips. This works great for larger marine species such as Chaetoceros, Thalassiosira species, Ditylum, etc. You can wash them by hand once or twice with sterile-filtered media, and unialgal cultures are easily established for many species (many people establish unialgal cultures successfully without even washing cells, and just re-isolate cultures that are not unialgal if necessary). You may or may not need to sieve... most often it isn't necessary.

Once the cultures are established, you can try different antibiotic dilutions. I would recommend first washing the cells with sterile media over a large pore-size filter (selected based on species size). An antibiotic mix that has worked well for me is:

Antibiotic mix 1:

1 mg/ml streptomycin stock

1 mg/ml ampicillin stock

I used these stocks to make the following antibiotic mixes in media, which I try in parallel:

500 ug/ml ampicillin, 100 ug/ml streptomycin

10 ug/ml ampicillin, 500 ug/ml streptomycin

50 ug/ml ampicillin, 50 ug/ml streptomycin

20 ug/ml ampicillin, 10 ug/ml streptomycin

10 ug/ml ampicillin, 5 ug/ml streptomycin

The idea is to find a high enough concentration that permits the target species to grow, killing off the maximum of co-occurring organisms. The Droop method is to do serial cross dilutions of the culture against an antibiotic mixture, to get different ratios of antibiotics to organisms. When selecting a mix of antibiotics to try, it is good to consider antibiotics that have different modes of action. You might find that one mix works well for some species, and others survive a different mix.

Often, it is suggested to add a small amount of peptone or other organic substrate (make a 1 g/L stock, and add to a final concentration of 1-10 mg/L). This stimulates bacterial growth, causing them to be killed by the antibiotics. I have done it successfully for different phytoplankton groups (diatoms, dinoflagellates, coccolithophorids) both with and without using peptone.

Treat for only 2-5 days, and then wash again or simply dilute into new media (depending on how fast-growing the species is). Let the treated cultures grow up once or twice before final testing. Test for bacteria by inoculating test cultures (50 ul per 10 ml) (media with 100 mg/L-1g/L peptone, for example) and by staining fixed samples with Sybr Green I, DAPI, or other fluorescent DNA stains to observe by epifluorescence microscopy. The ex-CCMP culture collection (NCMA) website https://ncma.bigelow.org has nice recipes for test culture media, which you can modify as needed. I imagine you could substitute many organic substrates for peptone (for example, yeast extract), but I haven't tried. Observe test cultures for up to one month to check for negative bacterial growth. If cultures are positive, repeat the procedure, but perhaps change the antibiotic mix.

Smaller diatoms, even planktonic ones, can be plated onto agar plates as suggested below, made with the same media you use to culture. Colonies will be visible between 2-3 weeks. The trick is to plate different volumes and dilutions, so you get nicely separated colonies. This will work well to isolate organisms like small Thalassiosiras or Cyclotellas, for example (I don't know how well it would work for raphid pennates with gliding motility). Usually, one would establish an initial culture on a plate made with media without antibiotics, then select the colonies one wants for growth in suspension, and then re-plate again on plates made with different dilutions of antibiotics, or you could treat the cultures in suspension with antibiotics, and plate them out. I haven't tried directly plating onto antibiotic-media, but I would be reluctant to do that because it takes time for the initial colonies to appear, and one wants to avoid selecting for antibiotic resistant bacteria (the idea is to keep antibiotic treatments short, so that if any do survive, they can be removed by a secondary treatment).

Another trick for the initial isolation is to use media that is 10- to 50-fold more dilute than normal. Most media recipes are very concentrated in nutrients, and those are not necessary, and high concentrations of nitrate or ammonium might even be stressful to some organisms. You might consider diluting your normal medium 1:10 with sterile-filtered water from the same sample from which you wish to culture.

There are many ways that will work and I suggest trying out several that are convenient for you and the organisms you wish to have. Don't feel you have to follow someone else's protocol either... think about your organism, how it behaves, and what might select for it over other organisms present. For example, perhaps you could use the gliding behaviour of benthic pennates to select them based on phototaxis. Or perhaps the organisms you are interested in will need more washing or less washing... don't be afraid to play and test a lot :)

I forgot to add one thing:

For testing for bacteria, both by culture and by epifluorescence, always use a positive and a negative control. The negative control is just sterile media. The positive control should be a non-axenic culture of the same species (preferably the same original strain) in a similar state of culture.

Please look at the detail review by Supriya and Ramachandra (2011)

http://wgbis.ces.iisc.ernet.in/energy/paper/marine_diatom_culturing/marine_diatom_culturing.pdf

If you can't get rid of the protozoans, try bubbling with N2-gas for an hour or two. Then isolate your target species, using one of the methods mentioned above, .

Hi Satya

I've never done it, but I think the best way to cultivate diatoms would be this: Petri dishes with agar medium containing the same water where you found the diatoms. Plate 1 ml of water with diatoms of your salmple very dilute. On the plate will form the various colonies, each with one species. Identify the species under the microscope. So from the colony mono-species takes a bit of diatoms and cultivate them in the same water where you found the diatoms plus specific mineral solution suitable for hydroponic cultivation of plants, but very diluted (1:20 to 1:50) (as for example Hoagland).

To control bacteria you must use antibiotics

I’m waiting for your results. Tell me them because I also would like to grow diatoms.

Best regards

Stefano

Hi Satya,

The best and fast way to isolate diatoms from a natural sample for a clonal and axenic culture is using a flow cytometer equipped with cell sorting (FACS). Using this technique you can get a 96 wells plate with theoretically 96 isolated cells in only a few minutes.

Best regards

Ivan

One of the ways is to use micropipette.If you don't have it, one of the method to make such a pipette is: hold a glass tube with its centre over a bunsen flame.As the glass melts,slowly & steadily pull both the ends of the tube in the opposite direction,till the middle portion comes out in a very thin streak.You now have 2 pipettes,but their ends are blocked by solid glass.Next break the fag end of one of the made pipette say a few millimetres each time,till you get the micro opening.See if that opening is just a little bigger than the size of the phytoplankton you want to culture.Then serially segregate the target item from the collected mixture from wild.

One way to get some amount of the starting material for culture is to first culture the segregated sample in a ' Goblet flask'.

Once you have the starting material,give optimum levels of photosynthesis for the culture, i.e.supply very small amounts of CO2 gas (which is the airation also),along with the required amount of Sunlight,+ the culture media.This will ensure faster growth of the target species.In case,if there is contamination by any other phytoplankton,then you will have to alter the physical & chemical inputs of the culture system.

Once the phytoplankton is harvested,place the slurry in a closed polythene bottle,remove out all the air,& then release CO2 gas over the water surface,so that there is some pressure of the gas in the bag.By doing this,all of the protozoans will get killed in a matter of a few hours - by exp. you have to quantify the time period.

In one of my field experiments,I used to use this method for removal of protozoans in a matter of 12 hours.But care needs to be taken to see that the bag with the phytoplankton slurry is placed in a fridge.

Isolation of microalgae dependson the type of microalgae. You can use capillary tubes under themicroscope for diatoms greater than 10 micrometres.For smaller ones it ismore convenient to isolate them in in TMRLmedium with 1.5% agar.

TMRL (Tungkang Marine Research Lab (Liao and Huang's Modified Medium)

KNO3 100.00 g

Na2HPO4.12H2O 10.00 g

FeCl3.6H2O 3.00 g

Na2SiO3.9H2O 1.00 g

Distilled water 1 liter

Utilization1 ml/L..

Please add the Na2HPO4.12H2O last to avoid precipitation.You may also add Penicillin G and streptomycin at 500UI/ml and 100ug/mL.respectively tocontrolcontamination.

Some important comments on the suggestion to use flow cytometry (FACS) to isolate diatoms. It's not necessarily as easy as a solution as hinted. I just placed an order for my own flow cytometer, and am sure going to love it when it comes, but it will not make the "old-fashioned" techniques obsolete for me, and I hope people starting out in microalgae/phytoplankton work don't get the idea that they need the "fancy expensive toys" in order to do great science. There are a lot of wonderful suggestions in this thread that rely on inexpensive materials, a good eye, and a bit of patience, and in many cases they will be superior to a flow cytometer. Here's why:

First, larger and chain-forming diatoms cannot efficiently be isolated by any commercially available cytometer, even from pure culture. Cytometers work best for cells less than about 20 um, although that can be pushed a little higher. Standard instruments cannot even measure those cells properly because they are larger... In normal instruments, the laser is focused to a point smaller than the size of those larger cells and the fluidics are not designed to accommodate large cells either (which can easily be isolated by hand anyway). I was part of a study that invented a special cytometer for measuring DNA content on Ditylum brightwellii (attached). Such instruments are not commercially available. If you try to run a culture of Chaetoceros, Thalassiosira rotula, Coscinodiscus or any species that is large (>30 µm) and/or chain-forming through a FACS Aria, you'll risk causing a major problem to the instrument, blocking up the fluidics and potentially putting the instrument out of order for days awaiting expensive maintenance.

For smaller cells, a cytometer can work great for helping purify cells, if you have access to one and can collaborate with someone who is very good at using one (you need someone with experience with diverse types of cells and who doesn't think of the instrument as just a black box where all you have to do is put the sample in and press a couple buttons). Most don't have ready access to a cytometer with sorting abilities, or it might be in some biomedical lab that only knows how to use it to sort lymphocytes and doesn't know that it's possible to change settings on one. I have used a FACS Aria to help me purify and make axenic Gephyrocapsa oceanica (coccolithophore) cultures (with help from Dominque Marie, a real expert in the use of these instruments), and found it was still necessary to incorporate antibiotics. The reason is fairly simple: Without special stains, the standard way to sort phytoplankton is based on detecting particles with chlorophyll fluorescence. Bacteria are not fluorescent, so the instrument won't see them and so cannot exclude them unless they are stained or if the instrument is seeing them based on scattered light.

From a natural sample, there is not an easy way to detect which cells are diatoms and which are something else (there is a stain that will be incorporated into the diatom frustule, and potentially could be used: see Le Blanc and Hutchins Limnol. Oceanogr.: Methods 3, 2005, 462–476). But normally a flow cytometer could not distinguish between a diatom and a flagellate (side scatter, or SSC, might help, but won't be perfect... See Olson et al. 1989 Cytometry 10: 636-643). A cytometer will allow an investigator to use 3-16 parameters to distinguish which cells to select (most use low numbers of parameters... it's honestly pretty tricky to keep track of >5 different parameters when selecting a population to sort). The human eye with a microscope can used a huge number of parameters to select which cells to isolate, if the cells are bigger than about 5-10 um (that's another of the many reasons cytometers are mostly used when investigators are targeting picoplankton, not microplankton and larger nanoplankton). However, I have made a few trials on natural samples with single-cell sorting on chl+ fluorescent cells of nanoplankton size characteristics, and a large portion of what can grow up was often small nanoplanktonic diatoms, which are then very clean clonal cultures (still not axenic if isolated by a FACS Aria). By the way, before putting natural samples in a flow cytometer, it's generally a good idea to pre-filter them through a 20-40 µm filter (to avoid problems).

That said, a cytometer can be used to help clean up a culture already established by "low tech" (or rather, human expert tech) methods, and it can work very well as long as the cells won't clog it.

As for instruments, the InFlux seems to be the best, because it makes droplets for sorting that are much smaller than those made by, for example, the FACS Aria and it can detect smaller particles and at a higher rate, both of which should allow for a more pure sort, but unfortunately even fewer people have access to an InFlux instrument. I have successfully established coccolithophore cultures that way from natural samples (that instrument can distinguish coccolithophores based on depolarization of scattered light). A FACS Aria can't do that particular trick, but it will work excellently for a lot of things you might want to sort (except large or chain-forming diatoms!), and I suspect it has a higher upper limit of what if can successfully sort than the InFlux (but I haven't tested). If you do try cytometry, make sure to trigger based on scattered light (SSC or FSC) and not chlorophyll fluorescence, and have the sensitivity high and threshold as low as possible, so that the instrument can be told to exclude all particles that are not desired. I have a lot more hints to offer, but I'll leave it at that.

The truth is, flow cytometers are not magic boxes that can solve our phycology problems without us doing any manual work, and they won't let us throw away all the "old fashioned" methods and tricks like the many offered in this thread by experienced investigators. Unless one really knows what one is doing or can work with someone who really understands a flow cytometer or one is willing to invest the time to develop such experience, such instruments won't help. Even if you do have the best instrument and know how to use it very well, you have to know what you are looking for and a cytometer is simply not going to work for larger or chain-forming species (which are a big fraction of the important diatoms). Finally, it takes more than an hour to get a cytometer properly set up for pure sorting. In that time, an experienced hand can isolate 20-50 cells of moderate to large-cell species manually using a microscope (I've seen some people who are even faster at isolating, doing about 1-2 cells/minute), just as many have suggested here.

Article Genome size differentiates co-occurring populations of the p...

I too agree with Norma about adding additional silicate into the F/2 medium!

the best method for diatom isolation that I found is dilution method and then use F/2 medium with silicate,

Serial dilution technique is the best method . For picking cells... you can use the IV -tube which you get in any medical stores, u can attach micro pipette tips at one end and the other end u can hold in your mouth for suction control. its better you use inverted microscope and as you keep going one level to other level of dilution ...each time change the front tip of tube with much narrow tip and at the final u can use glass capillary . important thing is that each time you need to keep washing tube or change the tube to avoid contamination . This will give you pure diatom culture but to make it bacteria free ... you need to use antibiotic treatment ... ( pls refer Patil and Anil 2005 MEPS, or D'Costa et al 2011)

Medium: f/2-medium (seawater, 0.2µm filtrated) add silicate (very important) and other nutrients according to Guillard and Ryther 1962, Canadian Journal of Microbiology,

Isolation: Using an aspirator tube (as described by Norma Zoe Neszi) and a glass capillary (as described by Chandravadan Trivedi) both you can get here http://www.krackeler.com/products/1106-Dispensers/15495-Calibrated-Micropipets.htm

Finally: You should try to establish different dilution rates in a 6er Cell Well plate (http://www.sigmaaldrich.com/catalog/product/sigma/z707910?lang=de®ion=DE) by a stepwise isolation of the diatom from the water sample and from this sample again and so on. This is a good way to get rid of other algae and detritus. To get the diatom bacteria free you have already some good suggestions.

Good luck,

Nils

Thank you all for your kind and valuable suggestions. - satya

Instead of subculturing after the initial pick with a pipette, you can dilute that pick in a drop of sterile water and then pick the cell of interest again, put it in a second drop of sterile water, et cetera untill all contaminating organisms are gone. This works best with an inverted microscope.

There is a long time that I did not performed such an exrecice. But when I did it I prevailed inexpensive material and a lot of patience.

I used to combine successive settling with decreasing times and sieving with weaker and weaker apertures. I added Na2HPO4.12H2O in the solution.