You will receive the culture strains in a screw-cap test tube. Slightly loosen the screw cap and keep the test tube in an appropriate place, as indicated in individual strain data. If you want to maintain the culture strain, please transfer the culture into fresh medium according to the following methods.
i) Before you receive the strains, prepare the appropriate medium according to the media list.
ii) Adapt the fresh medium to the culturing temperature.
iii) Transfer an appropriate quantity of cell suspension to the fresh medium by an aseptic technique. In the NIES-Collection, we transfer cell suspensions by using a sterilized pipette with a cotton plug (Photo 1. 2). Agitate the culture liquid by pipetting if cells settle out or become attached to the container when you are sucking up the cell suspension. In the case of cells such as those of Chattonella, which are weak and lack cell-coverings, gently suck up a concentrated part of the cell suspension without agitating to prevent breakage of the cells during the pipetting. The quantity of cell suspension differs with the species and the condition of the strain: to 10 mL fresh medium, we usually transfer 1 or 2 drops of cell suspension for small strains that grow well, whereas we use 4 or 5 drops for large strains and sparse cultures. In the case of agar slants, scratch a mass of cells off the surface of the agar with a sterile platinum loop and spread it on a fresh agar slant.
iv) Incubate the culture at the temperature and light conditions indicated in the individual strain data (Photo 3. 4). Transfer to new medium at the intervals indicated in the individual strain data (sometimes shorter or longer depends on your laboratory conditions).
v) In the NIES-Collection, we visually check the cultures every week and when needed with a microscope. If the culture does not grow well, we transfer again, and sometimes test other media and light conditions.
For heterotrophic strains, pay attention to the following points.
i) Some strains need cereal grain or other algae as food sources added to each medium during transfer (Photo 5). Others need algae multiplied in advance, in accordance with individual strain data.
ii) Incubation of these strains does not need light, except in the case of cultures that contain algae as food.
iii) Always agitate the culture liquid by pipetting before transferring. In the case of adherent strains, strong pipetting is needed.
You will receive several pieces of thallus. As soon as you receive them, transplant them into fresh culture media according to the following methods.
i) Prepare appropriate culture media before you receive the strains. Add 1 to 2 mL of germanium dioxide solution (1 mg/L) to a 900-mL glass vessel, each containing fresh medium. For unialgal strains, germanium solution is not necessary.
ii) Inoculate individual thalli gently into soil in a glass vessel by using a bamboo skewer or tweezers (Photo ii)-1, ii)-2 ). Make sure that one or more nodes of the thallus (root bulbils in the case of Lamprothamnium, stellate bulbils in the case of Nitellopsis) are embedded into the soil.
iii) Incubate transplanted cultures at the temperature and light conditions indicated in the strain data. About 2 weeks after the transplantation, the thalli should start to grow (Photo iii)-1, iii)-2 ). (You may place the cultures near a window in the laboratory, provided that the cultures are not exposed to direct sunlight or extremely high or low temperatures.)
iv) Transfer into new media at the intervals suggested in the strain data, by using the following methods.
a) Cut 3 or 4 apical internodes from a well-developed thallus with scissors or tweezers (Photo iv)-a）).
b) Remove microalgae from the surface of each piece with a paintbrush (Photo iv)-b）) and rinse with deionized water (or distilled water). (For unialgal strains this process is not necessary.)
c) Inoculate the rinsed pieces into a fresh medium as described in ii) to iii).
Media are generally composed of three types of components; macronutrients, trace metals, and vitamins. For convenience we recommend to prepare stock solutions of these components in dark glass bottles. Stock solutions of trace metals and vitamins are prepared at extremely low concentrations, and therefore required dilution steps. The following methods are currently used at the NIES-Collection.
Prepare stock solutions of individual macronutrients separately at a concentration of 10 mg/mL, and store them in a refrigerator (5℃).
These elements are prepared as either separate stock solutions or mixed stock solutions.
Prepare stock solutions of individual metals separately at concentrations of 1-10 mg/mL, and store in a refrigerator (5℃).
i) Prepare each metal solution as for the separate stock solutions shown in 188.8.131.52.
ii) Add approximately 80% of the final volume of distilled water in a beaker.
iii) First, dissolve the required amount of Na2EDTA, while stirring, if applicable.
iv) Add the required volume of each trace metal solution one at a time, while stirring.
v) Adjust to the final volume by adding distilled water, and store in a refrigerator (5℃).
Vitamins requirement is in majority fulfilled with three vitamins; vitamin B12, biotin, and thiamine HCl. Therefore, most of the media contain only these three vitamins. However, several media contain additional vitamins.
i) Prepare 0.1 mg/mL solutions of vitamin B12 and biotin and a 10 mg/mL solution of thiamine HCl. Disperse 1 mL of each solution into a separate micro-tube, and store in a freezer at -20℃.
ii) Thaw and dilute the vitamin solution to 1/100 to prepare stock solutions of 1 μg/mL vitamin B12 or biotin, and a stock solution of 100 μg/mL thiamine HCl. Store in a refrigerator (5℃).
Additional vitamins are added to some media as a mixture. We recommend to prepare a large volume of mixed stock solutions at once.
i) Prepare each vitamin solution at a concentration of 0.1-1.0 mg/mL. (Store these original solutions in a freezer at -20℃, if needed.)
ii) Add approximately 80% of the required volume of distilled water in a beaker.
iii) Add the required volume of each vitamin solution one at a time, while stirring.
vi) Adjust to the final volume by adding distilled water.
v) Dispense 100 mL of the vitamin mixture into several vessels, and store in a refrigerator (5℃) for use or in a freezer (-20℃) for storage.
Two categories of media are usually used; synthetic and enriched. The former is used for maintenance of all freshwater algal cultures and some marine ones and the latter for most marine ones. Most of the media are dispensed to test tubes and autoclaved before use, whereas some media must be filter sterilized.
i) Add approximately 80-90% of the required volume of distilled water to a beaker.
ii) Dissolve appropriate quantities of buffers such as Tris ( hydroxymethyl ) aminomethane ( known as Tris ), glycylglycine, HEPES, TAPS, Bicine, MES or 1, 2, 3, 4-cyclopentane tetracarboxylic acid ( if required ), while stirring.
iii) Add the appropriate nutrients from previously prepared stock solutions, while stirring.
iv) Adjust to the final volume by adding distilled water.
v) Check and adjust pH as specified in the media list with either 1 mol / L HCl or 1 mol / L NaOH ( if buffers are used ) or with either 0.1 mol / L HCl or 0.1 mol / L NaOH ( if no buffers are used ).
vi) Dispense 10 mL of medium into each test tube ( 18×150mm ) and sterilize by autoclaving ( 121℃, 20 min ).
i) Add approximately 80% of the required volume of distilled water to a beaker.
ii) Dissolve appropriate quantities of Tris, nitrilotriacetic acid (known as NTA) and major salts such as NaCl, MgSO4・7H2O, KCl and CaCl2・2H2O, while stirring.
iii) Add the other nutrients from previously prepared stock solutions.
iv) Adjust to the final volume by adding distilled water.
v) Check and adjust pH with 1mol/L HCl, if pH is specified in the media list. (usually pH 8.0)
vi) Dispense 10 mL of medium into each test tube and sterilize by autoclaving (121℃, 20min).
i) Collect offshore seawater free from pollution, and remove particulate matter by filtering through Whatman GF/F filters.
ii) Check salinity. (Usually salinity of offshore seawater is 35‰)
iii) Add approximately 80-90% of the required volume of seawater to a beaker.
iv) Dissolve appropriate quantities of Tris (if required).
v) Add the appropriate nutrients from previously prepared stock solutions.
vi) Adjust to the final volume by adding the filtered seawater.
vii) Check and adjust the pH to 8.0 with 1 mol/L HCl if required.
viii) Dispense 10 mL of medium into each test tube and sterilize by autoclaving (121℃, 20min).
MNK medium must be filter sterilized by using a filter apparatus with a filter (Millipore 0.22μm), which is previously autoclaved (121℃, 20min). Then, the medium is dispensed into previously sterilized test tubes by using a sterilized syringe or dispenser under aseptic conditions.
Agar is usually added at a concentration of 1.5% after liquid medium has been prepared, and before autoclaving.
i) Add the appropriate quantities of agar to the liquid medium and heat by autoclaving or on a hot plate.
ii) After melting, quickly dispense 10mL of agar medium into each test tube and sterilize by autoclaving (121℃, 20 min).
iii) After sterilization, lay the test tubes down with the upper part of the tubes elevated on a rod (1cm ø), and cool to form agar slants.
These media contain organic matter to encourage multiplication of bacteria as a food source for protozoa. For media containing wheat or rice grains, these cereals should be sterilized by dry heat (150℃, 30min) in advance, and kept in a cool place. For use, one grain of cereal is added to 10mL of medium.
Black soil, river sand, leaf mould, and garden lime used in the NIES-Collection are purchased from garden centers, whereas bottom mud from paddy fields, reservoirs, and ponds is collected by us. Soil quality influences the growth of Charales to a greater or lesser degree. Please refer to the media list and individual strain data for soil composition.
Freshwater strains: Deionized water (or distilled water). Brackish water strains: one-third to one-half diluted 1/3 Herbst ASW, i.e. the original medium is diluted to one-third to one-half with deionized water (or distilled water).
i) Put appropriate soil into a glass vessel up to one-quarter to one-fifth.
ii) Dampen the soil with deionized water (or distilled water).
iii) Cover the glass vessel with a plastic cap or aluminum foil, and autoclave it twice (121℃, 20min, overnight cooling down, and again 121℃, 20min).
iv) After the vessel has cooled down to room temperature, pour sterilized water (see 184.108.40.206 Water) into the glass vessel carefully (do not disturb the soil). When you make media for unialgal strains, use a clean bench (or a clean room) for this process.
Germanium dioxide solution especially discourages the growth of diatoms. To suppress the growth of undesired diatoms, add germanium dioxide solution (1mg/L GeO2) to the media.
i) Boil 200mL NaOH solution (1 mol/L).
ii) Add 0.5g GeO2 to the boiling NaOH solution very carefully．
iii) Cool down to room temperature.
iv) Check the pH and adjust to 7.8-8.0 with 1mol/L HCl.
v) Adjust to 500mL by adding deionized water (or distilled water).
vi) Autoclave (121℃, 20min).
Please click here to go to Media list.
A two-step freezing protocol is used in the NIES Collection: algal culture is cooled to -40℃ by a programmable freezer and then cooled rapidly to 196℃ in liquid nitrogen. Most cyanobacterial strains, some strains of green and red microalgae, and some strains of freshwater red algae are cryopreserved by the methods described in 4.1 and 4.2. Detailed methods for microalgae are also explained in Mori et al. (2002) and Mori (2007).
Mori, F., Erata, M. & Watanabe, M. M. 2002 Cryopreservation of cyanobacteria and green algae in the NIES-Collection. Microbiol. Cult. Coll. 18:45-55.
Mori, F. 2007 Cryopreservation methods of microalgae. Microbiol. Cult. Coll. 23:89-93. (In Japanese)
i) Culture: late log or early stationary phase cultures.
ii) Medium: appropriate sterile medium for each strain.
iii) Cryoprotectant: 6% dimethyl sulfoxide (DMSO) for cyanobacterial strains, and 10% DMSO for green and red algal strains dissolved in the appropriate media. These concentrations are double the final concentration. DMSO is previously sterilized by filtering through an alcohol-stable filter (Millex-LG).
iv) Laminar-flow cabinet and materials for aseptic treatment.
v) Cryovials: 2-mL presterilized polypropylene cryovials, pre-labeled with the strain number and date.
vi) Programmable freezer (e.g. Planer Kryo 320-1.7 is used in the NIES-Collection).
vii) Liquid nitrogen Dewar vessel: 10-L wide-neck Dewar vessel (Shattle Drum JIK-S10).
viii) Long forceps (19cm), cryogloves, a cryoapron, and goggles.
ix) Nunc polycarbonate storage boxes, 8-decker stainless-steel racks, a liquid nitrogen tank (Taiyo Nippon Sanso DR-245LM; vapor phase).
x) Water bath (e.g. As-One-Corp. Thermal Robo TR-1).
i) The processes ii)-iv) should be done under aseptic conditions.
ii) Dilute the cryoprotectant with appropriate medium to obtain double the final concentration, and cool it on ice.
iii) Dispense 0.5 mL of cell suspension (late log or early stationary phase culture) into each labeled 2-mL-cryovial.
iv) Add 0.5mL of the cryoprotectant (diluted and cooled) to each cryovial and mix well.
v) Leave the cryovials at room temperature for 15 min. vi) Place the cryovials in a programmable freezer (Photo 9), and start cooling at -1℃/min to -40℃.
vii) Hold the cryovials in the programmable freezer at -40℃ for 15 min.
viii) Transfer the cryovials rapidly to the Dewar vessel containing liquid nitrogen (Photo 10).
ix) After 1 h, transfer the cryovials in the Dewar vessel to a storage box and place the box on a stainless-steel rack set in the vapor phase of liquid nitrogen in a liquid nitrogen tank (Photo 11).
i) Preheat a water bath to 40℃.
ii) Shake the cryovials well in the water bath until the last ice crystal in the cryovials has melted (Photo 12).
iii) Under aseptic conditions transfer the contents of the cryovials into test tubes each containing fresh liquid medium. Incubate under dim light for a few days (depending on the strain), and transfer to ordinary culture conditions as suggested in the strain data.
i) Culture: several thalli cultured for at least 2 weeks after the last transplantation. If a thallus is large, cut it into small pieces with scissors or tweezers, and culture for more than 2 weeks (for recovery), before use.
ii) Medium: sterile Bold 3N medium.
iii) Cryoprotectant: 40% dimethyl sulfoxide (DMSO) for cryopreservation of Thorea okadae, T. hispida, and Nemalionopsis tortuosa; and 30% methanol for N. tortuosa. These concentrations are double of the final ones. DMSO and methanol are previously sterilized by filtration through an alcohol-stable filter (Millex-LG), and dissolved in sterile Bold 3N medium.
iv) Instruments: same as the instruments for microalgae.
i) Dilute the cryoprotectant (DMSO or methanol) with medium to obtain double the final concentrations (40% or 30%, respectively), and cool it on ice.
ii) Dispense a 0.8 mL aliquot of culture into each of the cryovials.
iii) Add 0.8 mL of 40% DMSO or 30% methanol to ii), and mix well. In the case of DMSO, leave the cryovials at room temperature for 15 min.
iv) Then same as 4.1.2 vi) to ix).
i) Preheat a water bath to 40℃, and cool appropriate amount of medium in ice water.
ii) Shake the cryovials well in the water bath, and transfer the cryovials into ice water just before the last ice crystals have begun melting.
iii) Transfer the contents of the cryovials quickly into 50-mL centrifuge tubes, add 40 mL of cold medium, and leave the tubes until the thalli have settled to the bottom.
iv) Remove the supernatant with a pipette.
v) Add 40 mL of cold medium again, and again remove the supernatant with a pipette after the thalli have settled.
vi) Transfer the thalli into 60 mL of fresh media in 100-mL conical flasks, and incubate under the culture conditions suggested in the strain data.
vii) All manipulations from iii) to vi) should be done under aseptic conditions.