Basic Neurospora protocols from the lab of D. E. A. Catcheside, Adelaide, Australia.




This manual documents common methods used for the culture of Neurospora in the Catcheside Laboratory at Flinders University. Last edited December 1999.





TITLE                                                                                                                            SECTION


PRECAUTIONS                                                                                                                          3




Tube cultures                                                                                                                                3

Liquid cultures                                                                                                                               3

Storage of cultures                                                                                                                        4

Growth tests to determine genotype                                                                                               4

Quantitative growth tests                                                                                                               5

Mating type tests                                                                                                                           5

Growth restriction techniques                                                                                                         6

Crossing methods                                                                                                                          6

Analysis of crosses                               Random spore analysis I                                                    6

                                                            Random spore analysis II                                                  7

                                                            Non-random spores                                                          8




Conidial Suspensions                I.          Small scale for subculturing                                                8

                                                II.         Medium scale for inoculation of multiple

                                                            tests or inoculation of liquid medium                                   9

                                                III.       Large scale conidial suspensions for

                                                            preparation of spheroplasts etc.                                         9

Mycelial fragments                                                                                                                        9



ISOLATION OF MUTANTS                                                                                                    10


RECOMBINATION ASSAYS                                                                                                  11




Vegetative growth:                    1.         Liquid Medium                                                                12

                                                2.         Slopes                                                                            13

                                                3.         Plates                                                                              13

                                                4.         Layer Agar                                                                     14


Crossing medium:                     1.         Tubes                                                                             14

                                                2.         Plates and Tubes




Vogel's X 50                                                                                                                               15


Trace elements                                                                                                                            15

SS X 20                                                                                                                                     16

SS (nit-2) x 20                                                                                                                            16

SGF X 20                                                                                                                                   16

Westergaard's X 4                                                                                                                      16

SC X 2                                                                                                                                       17

AWM X 4                                                                                                                                  17


SUPPLEMENTS                                                                                                                       18



                               CONVENTIONS USED FOR LABELLING MEDIUM:






Sucrose as C source

tube medium for stock cultures and tests, liquid medium for flasks

VM  +  supplements

(if any)

Sorbose with glucose and fructose as carbon sources

plating medium

SGF + supplements

(if any)

Sorbose with sucrose as carbon source

plating medium (less growth restriction)

SS + supplements

(if any)

Top layer agar with 1 M sorbitol

regeneration of spheroplasts

HARD TOP + supplements

(if any)

Top layer agar

plating of ascospores or conidia

TOP + supplements

(if any)




Neurospora has a reputation of being a troublesome air-borne laboratory contaminant. This is because the conidia are readily detached from their conidiophores.  Although there is no health risk, your experiments and those of others working with slower growing organisms could suffer from careless handling of Neurospora cultures.  Careful technique and a little thought will effectively prevent this potential problem and essentially all fungal contaminants will turn out to be those ubiquitously present in the air entering the laboratory; usually Penicillium or Aspergillus.  If you are unfortunate enough to create a cloud of conidia, cover all equipment then spray the laboratory air with a mixture composed of equal volumes of ethanol and 10% aqueous propylene glycol and retire from the room.  Having allowed time for the aerosol to settle, wipe all horizontal surfaces.  The spray will trap and settle airborne dust and spores but will create a sticky deposit on exposed surfaces. This will evaporate eventually but may damage absorbent materials that are not covered.



Tube cultures


Neurospora is conveniently cultured on 1.5 ml agar slopes of Vogel's N minimal medium (supplemented with specific growth factors as required) in 100 x 13 mm tubes plugged with non-absorbent cotton wool.  Such cultures are sufficiently small to handle in large numbers but provide sufficient conidia for initiating most types of experiment.  Growth is most rapid at 34o but stocks containing the colonial temperature sensitive mutation cot-1 must be grown at 25o to obtain conidia.  Conidiation can be improved by moving the tubes into diffuse light at room temperature after conidiation has started (i.e. after 18-24 hr. at 34o, 40-48 hr. at 25o).


Large numbers of conidia can be produced from a 100 ml layer of agar-solidified Vogel's medium in a 1 l conical flask.



Liquid cultures


Submerged cultures of laboratory strains do not conidiate.  Liquid Vogel's N medium is used.  The yield of mycelium is about 30 g fresh weight/l.


Aeration may be achieved by shaking; use flasks plugged with cotton or closed with loose fitting metal caps, containing 1/5 the flask volume of medium, and adjust the shaker to give vigorous mixing without splashing the plug.  With care, up to half a flask volume of medium can be used; particularly on orbital shakers but under aeration is a common problem when this is tried and poor growth is likely.  At 25o growth is complete in about 72 hours.


Larger vessels may be aerated by sparging with compressed air sterilised by passage through a tube loosely packed with non absorbent cotton wool.  The glass sparge tube should be inserted deep into the medium and the air flow adjusted to give good vertical mixing.  8 l of medium in a 10 l capacity bottle is satisfactory.



Storage of cultures





Cultures grown on slopes, suitably supplemented, can be kept at 5o following incubation and will normally remain viable for several months.  At -20o, the tubes remain viable for several years.



Long term


The most convenient method for long term storage is on silica-gel.  Half fill 100 x 13 mm tubes with 12-20 mesh indicator free silica-gel granules, plug with rolled muslin, cover plugs with foil and sterilise at 160oC for one and a half hours in an oven.  Store sterile tubes over anhydrous calcium chloride in a desiccator.  Dispense 1.5 ml of reconstituted dried skim-milk per 100 m x 13 mm tube, cap and autoclave 5 min. at 121oC.  Store the tubes at room temperature and on each of next two days steam the tubes for half an hour.  Prepare 1.5 ml tubes of sterile water and use these to add 1.5 ml of sterile water to a 3-7 day old conidiating culture, suspend conidia and then add 1.5 ml sterile skim milk.  Mix and allow the conidia to settle out ( 0.5 hr).  Don't be tempted to try and suspend the conidia directly in skim milk, it is very difficult. When the conidia are settled, use a pasteur pipette to remove about half of the supernatant and resuspend the conidia in the remainder.  Before opening, hold each gel tube horizontal and shake the gel to provide an air gap to the base of the tube.  Then using a pasteur pipette, distribute about 0.8 ml of suspension evenly over the granules in each of the two silica gel tubes.  Shake the gel periodically until all of liquid is absorbed then dry the tubes in vacuo over CaCl2 for 2-3 days.  Cover the tubes with parafilm and store dry at 5o.  Preparations made in this way remain viable for many years.  Several hundred subcultures may be made from each tube.  To subculture, tip a few silica gel granules onto a slope of appropriate medium and incubate.  Reseal the gel tube and return to 5o.


Aconidiate strains can be preserved on silica gel.  Grow in tubes of liquid medium containing sterile glass beads.  Vortex mix after growth and add the resulting broken mycelium to skim milk.  Treat as for conidia.



Growth tests to determine genotype


If a culture contains an auxotrophic mutation it will not grow properly, if at all, in the absence of its specific growth requirement.  Other mutations confer resistance to antimetabolites and enable growth on media toxic to the wild-type.  Tests for ability to grow on Vogel's minimal medium supplemented with different combinations of growth factors enable determination of the genotype of multiple mutants.  A control medium on which all mutant combinations can grow should always be used.  Incubation at different temperatures enables the identification of temperature sensitive mutants.  Tests may be performed in tubes or on plates ruled into a grid of 16, 25 or any other convenient number of squares.  Test media may be inoculated with conidia or small pieces of mycelium cut from a colony growing on an agar plate.  When plates are used, a technique to restrict growth is essential (q.v.).


Quantitative growth tests


Use flasks of liquid medium supplemented as required.  Harvest over a buchner after growth is complete (or earlier if plotting a growth curve); wash with distilled water, wrap in weighed foil and dry at 60-80oC to constant weight.  Flasks can be incubated without shaking but more reliable results are obtained by aerating the flasks by shaking them on a reciprocating or orbital shaker.  For reciprocating shaker, fill the flasks with 1/5 volume of medium.  For orbital shakers, somewhat larger volumes can be used.  Shaking speeds should be adjusted to give vigorous aeration without splashing the plugs; the precise speed will depend upon the geometry of the flask.  Cotton plugs should not be too tightly rolled and metal thimble caps are a good substitute since they permit good gas exchange.  Flasks with uneven neck size or which are unevenly plugged, may give variable growth yields.



Mating type tests


(i)         By using standard crossing methods (q.v.) and pairing the unknown with each of two standard strains, one A, the other a mating type,




(ii)        Inoculate the centre of a petri dish of Westergaard or SC agar medium with the fluffy a mating type tester and a second dish with a fluffy A tester.  The fluffy mutant is aconidiate and thus safe to grow in petri dishes.  Normal strains will grow out of the plate, conidiate, and cause contamination problems. Incubate for 5 days at 25o, check for the present of protoperithecia, rule mycelial mats into 16-25 squares with a sterile needle and test unknowns by inoculating one square on each mating type tester with a dab or conidia.  A single loopful of conidia can be used for both plates since there is little danger of transferring the aconidiate tester from one plate to the other.  Incubate at 25o and score for perithecial development at 18-48 hr.


(iii)       Inoculate 1.5 ml slopes of SC medium in 100 x 13 mm tubes with fluffy testers.  Incubate 5 days at 25o, check for perithecia and store at 4o for up to 4-6 weeks before use.  Inoculate with unknown, incubate at 25o for 24-48 hr.



Growth restriction techniques


Sorbose is not utilised by Neurospora, it causes rupture of hyphal tips and stimulates branching growth.  Two sugar mixtures are commonly used in place of sucrose, SGF being the most restrictive.


            SGF:    1% sorbose, 0.05% glucose, 0.05% fructose

            SS:       1% sorbose, 0.1% sucrose.


These are made as 20 x stock solutions, sterilised by autoclaving, stored at 4o and added aseptically to pre-sterilised medium.  Colonies will be normally be visible after 24 hours and well grown by 48 - 72 hours.  Plates should be disposed of or transferred to 4o after this time or conidiation may occur.


The colonial temperature sensitive mutant cot-1 C102t gives tightly restricted dense growth at 34o.  Incubation at this temperature immediately following plating may prevent "visible" colonies developing for some days, however, very large numbers of colonies may be screened on each plate.  It is convenient to incubate cot-1 conidia or ascospores for 18-24 hr at 25o on SS or SGF medium then transfer to 34o for 24-48 hr.  This yields star like colonies which are easily visible to the unaided eye.



Crossing methods


Crosses are conveniently performed in 150 x 16 mm cotton plugged test tubes containing 4 ml of liquid Westergaard's medium supplemented as necessary and a 6 cm x 4 cm spill of Whatman #1 filter paper folded lengthwise into a "W".  Normally conidia from both parents are added together, mixed and spread evenly over the paper by inclining and rotating the tube.  If it is desirable to have only one parent acting as female, inoculate the female parent first, incubate for  5 days or until protoperithecia are present, then wet the culture surface with a conidial suspension prepared from the male parent; decant excess suspension.  Crosses should be incubated at 25o.  Ascospores are normally produced in 5-20 days depending upon the cross.  If genetic analysis is intended, wait  7 days after the first spores are discharged from the perithecia to ensure full maturation of the different genotypes.  Maturation of spores can be speeded by incubation at 30o following their discharge but this will prevent further meiosis.





Ascospores must be heat treated to induce germination.  1 hour in a 60o water bath is more than adequate and serves to kill conidia.  The heat treatment should not be interrupted as ascospores will start to germinate within minutes and are killed by further heating.  The heat treatment can usually be shortened to 30 minutes or prolonged up to 2-3 hours without serious loss of ascospore viability.  It is usually convenient to heat shock spores in suspension prior to layering onto plates.  If spores are plated prior to the heat shock, use a 60o incubator and allow additional time for the plates to warm-up.


Random spore analysis I


With a loop of sterile water, transfer about 50-100 ascospores into a drop of sterile water on a suitably supplemented plate of SGF or SS medium and spread the spores with brisk strokes: rotate the plate at the same time to obtain an even distribution.

Text Box:

To make a spreader, seal the end of a pasteur pipette in a bunsen flame and bend it into shape or make one from glass rod.  Sterilise by dipping in alcohol and flaming, but don't hold the spreader in the flame.


More efficient spreading can be achieved by making a spore suspension in soft agar and layering 3 ml on each plate.


Incubate the plate for 60 min at 60o and transfer to 25o for 24 hours or until colonies are visible.  After checking under the dissecting microscope that the colony is derived from the growth of a single ascospore, pick a small portion onto a slope of suitable medium, grow to conidial stage and test genotype (q.v.).  Alternatively, pick small portions of each colony onto plates of test medium.


This method has two disadvantages: the proportion of viable ascospores is not determined and truly random isolation of the colonies produced by germinated spores is difficult.  There may be an unconscious operator effect preferentially picking colonies having a particular appearance; an appearance often dictated by genotype.  Hence genetic ratios determined by this method may be biased.



Random spore analysis II




Text Box:  Cut a 1 cm x 3 cm block of agar from a plate and divide each of the longer edges with  3 mm cuts (razor blade or microspatula) into about 25 "keys".  Streak a loopful of spores down the centre of the block then using a loop made from fine human hair (double a strand of hair into a capillary drawn on the end of a pasteur pipette and fix it in place with epoxy glue), pull one black spore onto each key.  Take care not to lose spores in the cuts or mixed cultures will result.  Colourless spores will be inviable, make note if their frequency exceeds the normal 1-2%.  Do not attempt to sterilise the loop and work under a comfortable magnification (around 25x).  When all keys have a spore, cut under each key (I) and behind each spore (II).  Using a spade ended needle, transfer each block to a slope of suitably supplemented medium.  Take care that the spore is not lost during the transfer.  Heat treat slopes at 60o in a water bath for one hour and incubate at 25o until conidia are produced then carry out tests to determine the genotype.

With practice this method is as fast as the first, all isolation manipulations are performed on one day instead of two and the number of tubes in which no growth occurs measures the proportion of inviable spores.  Such inviability may be associated with particular genes or gene combinations.  A variation on this method is to spread ascospores onto an agar plate then pick black spores singly into tubes using a spade ended needle.


Non-random spores


It may be desirable to select particular genotypes in which case method I can be employed and selective medium used for plating the spores. Rare genotypes can often be obtained with relative ease by germinating spores top layered onto a selective medium, marking all those that grow and then overlaying the plate with a second top layer that contains an additional nutrient and incubating again. Given an appropriate choice of medium, the colonies that grow after the second incubation will be enriched for the genotype you need.




Always work near a flame and in a draft free area.  Mop the bench with a disinfectant prior to starting work.  (Absolute alcohol is suitable but beware of flames).  When handling cultures it is desirable to flame the neck of the vessel immediately on opening and again immediately before reclosing.  This may be impracticable with certain large scale cultures and with petri dishes but is imperative with stock solutions where contamination poses serious problems.  When opening cultures, keep hold of the plugs or caps, don't place them on the bench.




Take care not to disperse conidia into the air.  The yield of conidia from a culture can usually be improved by removing from the incubator into normal room light about 24-48 hours after conidiation commences.  The viability of conidia will start to decrease after  7-10 days storage at room temperature.  This period can be prolonged by storage at 4o.


I.          Small scale for making subcultures.




Flame a wire loop.  When doing this, hold the loop nearly vertical in the flame to get whole length of wire red hot.  Cool the wire in a petri-dish of sterile water and pick up a generous loopful of water.  Open the culture, flame neck of tube and holding tube and loop horizontal, touch the loop on the wall of the tube near to the conidia to deposit a droplet, touch the loop lightly onto the conidial mass, then puddle in the droplet to suspend conidia.  Pick up a loopful of the suspension, tap wire lightly onto the wall of the tube to remove any dry conidia, withdraw the loop from the tube and hold it stationary whilst reflaming neck of culture tube and replacing the plug.  The loop may now be used for inoculating one to several tubes or marked squares on plates.  Flame the neck of the subculture tube on opening and reclosing.  It is possible to hold both the culture and subculture tube at the same time in one hand and manipulate both plugs and the inoculating loop with the other hand.


With care, a single loop of conidial suspension can be used to place conidial dabs in 2-3 tubes or plates thus saving labour when testing nutritional mutants and mating types.


II.         Medium scale for inoculation of multiple tests or inoculation of liquid media.


Open the culture and a tube containing 1 ml of sterile water, hold the necks of the tubes inside a "cold" (luminous) bunsen flame and tip into the culture.  This should be done carefully but quickly to avoid overheating the water.  Hold tube necks inside flame for a few seconds to cremate any escaping conidia that have been displaced by the addition of water to the tube and return the plug to the culture tube as it is withdrawn from the flame.  Leave the tube for a short time to avoid scorched fingers, then vortex mix to produce a conidial suspension.  Leave tube again for a short time to allow any airborne conidia in the tube to settle and then remix.


To inoculate large numbers of differential test media, use a sterile pasteur pipette as a capillary (no rubber teat) and touch onto the appropriate squares on test plates.  The pasteur will pick-up 50 l or more if dipped into the conidial suspension and will deliver small inocula when touched onto an agar surface.


To inoculate liquid medium, use a pasteur pipette and rubber teat.  1 drop per 50-100 ml is adequate.  The whole 1.5 ml suspension is adequate for a 8 l carboy culture.


Note that supplements present in the slope culture will be leached out into the suspension.  This is not normally a serious problem but for critical work the suspension should be removed from the culture tube as soon as possible. Even better, pellet the conidia by centrifugation and resuspend them in a fresh aliquot of sterile water.


Note also that the suspension will contain some mycelial fragments.  If it is imperative to remove these, filter the suspension through two layers of sterile muslin held in a filter funnel or in the neck of a flask or over the mouth of a pasteur pipette as a small sock held in place with autoclave tape.


III.       Large scale conidial suspensions for preparation of spheroplasts etc.


Inoculate 1 l flasks containing 100 ml of suitably supplemented Vogel's medium solidified with agar and harvest the conidia by suspending them in 20 ml sterile water.  Filtration through muslin and washing by centrifugation is usually necessary. Muslin filters are conveniently made by folding single or double layers into a cone (as for filter paper) and taping into a funnel. This can be wrapped in foil or paper prior to sterilising.




Aconidial cultures must be transferred as mycelium.  A flamed needle or spade, cooled in air or in sterile water, is satisfactory for transferring a few strands of the aerial mycelium to a fresh slope.


Colonies growing on restrictive media in petri dishes are subcultured by cutting out small pieces of agar containing some mycelium, using a spade ended needle.  A small inoculum is particularly important when transferring from sorbose agar or the sorbose will inhibit growth in the new culture!  It is convenient to use two spade needles in rotation, the needle not in use being stood vertically in a holder to cool after being flamed to red heat.




Mutants resistant to antimetabolites, (e.g. analogous of metabolites) suppressor mutations, or reverse mutations to prototrophy can be selected directly by plating conidia treated with a mutagen onto a suitable selective medium.  Sorbose or cot-1 is used to restrict growth.  Auxotrophic mutations can be selected by filtration enrichment.  Mutagen treated conidia are cultured on liquid Vogel's medium supplemented (if required) to permit growth of non-mutant conidia.  Non-mutant conidia are able to grow and the resulting colonies can be filtered out of the suspension culture with sterile muslin or absorbent cotton wool.  When all growth has ceased, the remaining ungerminated conidia are pelleted by centrifugation and plated on medium supplemented with the specific growth requirement of the mutant sought and grown under growth restricting conditions using cot-1 or sorbose.  Colonies which grow are then tested for the specific growth requirement.  50% or more of the survivors of the filtration may be mutants of the desired type, though 5% or less is more normal.




Suspend conidia from a 1.5 ml slope culture in 20 ml sterile water, filter aseptically through a double layer of muslin, pour into a petri dish, set shaking, remove lid and irradiate to  95-99% kill with ultra-violet light.  Since the appropriate dose is likely to vary from strain to strain, an experiment to determine the necessary doe will be needed first.  A convenient way of shaking the petri dish is to clip it to a vortex mixer and adjust the vibration speed to produce a standing wave.  Pipette the irradiated suspension into 100 ml of Vogel's minimal (or another appropriate medium) containing 2% sucrose.  Incubate with shaking at 25o, or at 34o if temperature-sensitive mutants are sought.  Refilter as often as necessary until there has been essentially no growth for 24 hours.  (It is normally necessary to filter at 8-12 hour intervals but heavy growth in the period 18-30 hours may require filtrations more frequently to prevent ungerminated conidia from entangling in and fusing with mycelial growth.  When growth has more or less ceased, refilter and plate the remaining ungerminated conidia onto supplemented medium.  If cot-1 has been used, this can be done by heating the suspension to 45o adding an equal volume of 45o molten medium containing double strength agar, sorbose and any supplements.  Pour the mixture into plates allow to solidify, incubate overnight at 25o and then transfer to 34o for further incubation.  If cot-1 has not been used, the sucrose in the medium must be removed.  To do this add a suspension of the conidia from a 1.5 ml slope, which have been killed by incubating the tube at 60o for several hours, and pellet the conidia by centrifugation.  Resuspend the conidia in a small volume of sterile water, add to molten supplemented SGF medium, plate and incubate at 25o.  Following a suitable period for growth, pick colonies of putative mutants into tube of fully supplemented medium prior to testing for auxotrophy.  Alternatively, transfer small inocula to unsupplemented plates and look for slow growth.  Re-pick the slow growing colonies back onto supplemented medium, grow to conidia and then test fully.





When analysing crosses between closely linked genes or alleles of the same gene, measurement of recombination frequency normally demands selective plating techniques in which large numbers of ascospores are plated onto a medium which permits the growth of one of the two recombinant classes, the prototrophic recombinants.  The number of recombinants may be related either to the total number of ascospores plated or more conveniently to the total number of viable ascospores.  The former can be estimated by counting the number of ungerminated and germinated spores on an area of the selective plate and multiplying up to obtain the total count.  Viable spores are estimated by plating out a suitable dilution onto a non-selective medium on which both the recombinant and parental types can grow and multiplying by the dilution factor.  Frequently, different numbers of selective and non-selective plates are used and you will need to take this into account when calculating the prototroph frequencies.  A convention of this laboratory to express recombination frequency in terms of map units (% recombination) when considering non-allelic recombination and in terms of prototrophs per 105 viable ascospores when considering recombination between alleles of the same gene.  The following method is suitable for estimating prototroph frequencies of between 1 and  1000/105.  For each analysis, the following are required:


            .           A mature cross

            .           A 28 x 200 mm sterile tube

            .           A bottle containing 100 ml sterile water

            .           two 100 ml winchesters containing 19 ml minimal layer agar held at 60o

            .           one 100 ml winchester containing 19.5 ml minimal layer agar held at 60o

            .           5 selective plates

            .           3 non-selective plates.


Taking care not to disperse the conidia, add  8 ml of sterile distilled water to the crossing tube.  Vortex mix briefly and pour the spore suspension through a muslin filter which will retain perithecia, mycelial fragments and fragments of filter paper.  NOTE:  over zealous mixing will disrupt the filter paper.  The fragments may then prove a nuisance in later stages of the analysis.  Add a further  8 ml of sterile water to the crossing tube, re-mix and filter.  Repeat if necessary to wash the filter paper onto the muslin.  Rinse the muslin with the remaining sterile water, collecting all of the washings into the 28 x 200 mm tube.  A sterile spatula is useful for retrieving the paper and perithecia from the cross tube and for pressing the filter to dislodge spores. Rinse the spatula in water between crosses then dip in alcohol and flame sterilise. A water rinse is needed as ascospores remain viable in ethanol for some time. Ensure the rinsings end up in disinfectant.


Allow the tube to stand for 1-3 hours away from any source of heat which may cause convection.  The spores will settle out to form a black layer on the bottom of the tube.  Decant the supernatant gently into disinfectant, leaving about 2 ml of ascospore suspension.  Swirl the spore suspension and pour into 19 ml of minimal layer agar held at 60o, this is Bottle A.  Incubate for 60 minutes to kill conidia.  For the remaining manipulations, warm the pipettes by flaming prior to use and return the bottles to 60o as soon as possible.  Also, speed is of the essence since the spore will settle-out rapidly: mixing must immediately precede each pipetting operation   Vortex mix, pipette 1 ml into the second bottle containing 19 ml of molten minimal layer agar, bottle B 1/20 dilution.  Vortex mix bottle B and pipette 0.5 ml into 19.5 ml of molten minimal layer agar, bottle C 1/800 dilution.  Vortex mix bottle C pipette 3 ml onto each of 3 fully supplemented plates, "C" plates.  In cool weather it may be helpful to prewarm the plates to 60o to aid spreading.  Vortex mix bottle A and pipette 3 ml onto each of 5 selective plates, A plates.  For recombination frequencies in the high range, or if the frequency range is unknown, also plate from the B bottle onto selective medium.


In most instances, incubate the plates at 25o for  18 hours and transfer to 34o for 24 hours.  Count colonies and calculate prototroph frequencies.  When discarding plates, ensure that they are not left at room temperature or growth will escape from the plates in a few hours.  Keep discard plates overnight in the 60o incubator or autoclave them the same day.


When analysing recombination between nit-2 alleles, incubate A plates for 42 hours at 25o and then 48 hr at 34o.


This method employs 2% sucrose in the layer agar with the plates containing SGF plates. Excepting in analyses of nit-2 crosses where SS is used in the A plates.  Both parents must be cot-1.  Non cot-1 parents require SGF layer agar and lower dilutions may have to be used since the colony size will be large.  If spore yields are high or low, it may be desirable to vary the dilutions to give 1/200 to 1/1600 in the C bottle.  When very low recombination frequencies are being measured, the spores from two or more crossing tube may be combined at the filtration stage.  In this case, reversion rates may become significant and it is desirable to set up control isoallelic crosses.







1.         Liquid medium


            Vogel's 50 X stock                                 2 ml

            Sucrose                                                  2 g

            Supplements                                             as needed

            Distilled water                                       100 ml


            (pH should be 5.8)


            Autoclave: 10 psi for 10 min. is satisfactory for small volumes (say to 200 ml) but for larger volumes 15 psi for 15 min. may be needed. Give 8 l carboys at least 20 min.



2.         Slopes    (make 100 ml per basket of 13 x 100 mm tubes)


            Vogel's 50 X stock                                 2 ml

            Sucrose                                                  2 g

            Supplements                                            as needed

            Difco Agar                                             2 g     Note that it is useful to mix the sucrose and agar dry prior to adding liquids. This helps disperse the agar and reduces problems with lumps and proper mixing.

            Distilled water                                       100 ml

            (pH should be 5.8)


            Steam, or autoclave 5 min to dissolve agar.  Mix well and dispense 1.5 ml per tube, taking care not to get agar where it will wet the plug or the plugs will stick. Plug the tubes with non absorbent cotton. A push plug formed by inserting a tuft of cotton with forceps is satisfactory.  Autoclave at 10 psi for 10 min (longer if the autoclave load is substantial).  Slope the tubes whilst they cool. The agar should not be more than about half way to the plug or the mycelium is liable to grow into the plug.



3.         Plates   (allow 20 ml/plate)


            Vogel's 50 X stock                                 2 ml

            Supplements                                             as needed

            Difco agar                                              2 g

            Distilled water                                       100 ml


            (pH should be 5.8)


            Autoclave 10 psi 10 min or more as appropriate, then add aseptically:


            SGF or SS 20 X stock                         5 ml



            Mix, cool to less than 50oC and pour into petri dishes.  Flame surface to break bubbles before agar sets.



4.         Layer agar        (Make on day of use)


            Vogel's 50 X stock                                 2 ml

            Sucrose                                                  2 g     only for analysis with cot where plates are incubated at 25o then moved to 34o

            Difco agar                                            0.8 g

            Supplements                                         as necessary *

            Distilled water                                       100 ml



            Autoclave 10 psi 10 min or 15 psi for larger volumes.  Dispense and keep at 60o in a water bath.  * Normally supplements are not added to layer agar.




1.         Tubes


            EITHER           Westergaard 4 X stock             25 ml

                                    Sucrose                                    2 g

                                    Supplements                             as needed

                                    Distilled water                           75 ml


                                    Adjust pH to 6.5.



            OR                   SC Medium 2 X stock              50 ml

                                    Sucrose                                    2 g

                                    Supplements                             as needed

                                    Distilled water                           50 ml


                                    (pH should be 6.5 without adjustment)


            OR                   For crosses between nit-2 mutants, use AWM 4 X in place of Westergaard's.


            Dispense 4 ml/tube, containing 6 x 4 cm spill of Whatman # 1 filter paper folded lengthwise into W.  Plug and autoclave 10 psi 10 min or 15 psi 15 min for larger volumes.


2.         Plates and tubes for mating type tests


            As above but with 2 g Difco agar per 100 ml.  Autoclave and dispense 20 ml per plate or 1.5 ml per tube, plug and slope.





VOGEL'S MEDIUM N, 50 X STOCK SOLUTION (to make 1 litre)


Dissolve in the order listed.  Do not add any component until the previous one has dissolved completely.  Use a magnetic stirrer.  Gentle heat may be used.  BDH "analar" chemicals are recommended.


            Distilled water                                                   700 ml

            Na3 citrate.  5 H20                                         150 g (.2H20, 130 g)

            KH2PO4 anhydrous                                          250 g

            NH4NO3 anhydrous                                         100 g

            MgSO4 . 7H20                                                 10 g

            CaCl2 . 2H20                                                      5 g     dissolved in 50 ml distilled water

            Trace element solution                                        5 ml

            Biotin                                                               25mg

            Chloroform                                                         2 ml (as a preservative)

            (this will be pH 5.8 when diluted. Do not adjust pH of stock)                


            Store at room temperature in a dark bottle.  Check for presence of preservative from time to time.





Dissolve in the order listed.  Use BDH "analar" chemicals where possible.


            Distilled water                                                   95 ml

            Citric acid 1H20                                                5 g

            ZnSO4.7H20                                                     5 g

            Fe(NH4)2(SO4)2.6H20                                      1 g

            CuSO4.5H20                                                    0.25 g

            MnSO4.1H20                                                   0.05 g

            H3BO3 anhyd                                                   0.05 g

            Na2MoO4 . 2H20                                             0.05 g

            Chloroform                                                       1 ml (as a preservative)

            Store refrigerated in a dark bottle.






            Sorbose                                                           20 g

            Sucrose                                                              1 g

            Distilled water                                                   100 ml


            Autoclave 10 psi 10 min.  Store refrigerated.



SS (nit-2) 20 X STOCK SOLUTION for use when plating cross homozygous nit-2 and am-1.


            Sorbose                                                           10 g

            Sucrose                                                              2 g

            Distilled water                                                   100 ml


            Autoclave 10 psi, 10 min.  Store refrigerated.





            Sorbose                                                           20 g

            Glucose                                                              0.5 g

            Fructose                                                             0.5 g

            Distilled water                                                   100 ml


            Autoclave 10 psi, 10 min.  Store refrigerated.





            KNO3                                                              4 g

            KH2PO4                                                           4 g

            MgSO4 . 7H20                                                 4.1 g

            CaCl2 . 6H20                                                    0.8 g (.2H20, 0.54 g)

            NaCl                                                                0.4 g

            Trace element solution                                      0.4 ml

            Biotin                                                                10 mg

            Distilled water                                                   1 litre

            Chloroform                                                       2 ml (as a preservative)


            Store at room temperature in a dark bottle.  Check for presence of preservative from time to time.




            KNO3                                                              2.0 g

            K2HPO4                                                           1.4 g

            KH2PO4                                                           1.0 g

            MgSO4 . 7H20                                                  2.0 g

            NaCl                                                                0.2 g

            CaCl2                                                               0.2 g

            Biotin                                                               10 mg

            Trace element solution                                      0.2 ml

            Distilled water                                                   1 l

            Chloroform                                                       2 ml


            Store refrigerated in a dark bottle.



AWM 4 X STOCK SOLUTION for use in crosses homozygous nit-2.


            NH4NO3                                                          2 g

            KH2 PO4                                                          4 g

            MgSO4 . 7H20                                                 4.1 g

            CaCl2 . 6H20                                                    0.8 g (.2H20, 0.54 g)

            NaCl                                                                0.4 g

            Trace element solution                                      0.4 g

            Biotin                                                                10mg

            Distilled water                                                   1 litre

            Chloroform                                                       2 ml (as a preservative)


            Store in a dark bottle at room temperature.  Shelf life beyond 1 month not proven.


SUPPLEMENTS                                                                                 stock solutions*


                                                mg/100 ml medium       mg/100 ml        drops/1.5 ml slope


adenine                                                20

adenine sulphate                                   30

adenosine**                                        40                        500                         1

L-alanine                                             50                        400                         5

p-aminobenzoic acid                              0.2                        3                           2

anthranilic acid                                     13

arginine                                                50

asparagine                                           30                        400                         1

(in presence of alanine)                          20

choline chloride                                    38

L-cysteine HCl                                    20

glycine                                                 150

L-histidine                                            20                        800                         1

L-homoserine                                      20

indole                                                    2

inositol                                                   5                         100                         1

L-isoleucine                                         23

L-leucine                                             20

L-lysine                                               40                        600                         2

L-methionine                                        50

monosodium glutamate                         40

nicotinamide                                           1

D-pantothenate                                      1

L-phenylalanine                                    20

L-proline                                             50

pyridoxin HCl                                        1

L-serine                                               20                        300                         2

sulphanilamide                                        3.5

thiamine (aneurin)                                   1

L-threonine                                          10

L-tryptophan                                       40

L-tyrosine                                            20

uracil                                                   10

uridine                                                 10                        150                         2

L-valine                                               20


*          autoclave and refrigerate


**        best for vegetative growth; adenine is better for crosses.




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