1.2 Storage & Re-isolation of frozen cultures 5

1.3 Testing the phenotype of a strain 6

1. ploidy 6

2. mating type 6

3. testing the genotype by PCR 7

4. temperature/ cold sensitivity 8

5. auxotrophy 8

6. adenine mutants 9

1.4 Growing cells

generation time 9

liquid cultures 10

1.5 Fission Yeast Classical Genetic Techniques

genetic crosses 11

genetic analysis of products of meiosis 12

tests for allelism 13

construction of double mutants 13

tests for dominance/recessiveness 14

isolation of diploid strains 15

protoplast fusion 17

protoplast fusion2 18

1.6 Mapping 17

1.7 Mutagenesis of yeast strains 19

1.8 Table of media ingredients 21

S. pombe markers 23

Autonomous replication sequences 23

Expression vectors 24

Promoters 25

2.2 Transformations

Lithium Acetate methods, I and II 26

Electroporation 28

Protoplasting 28

Lithium Chloride 30

2.3 Integration of a plasmid into the genome. 30

2.4 Gene Disruption and replacement 31

2.5. Stability 33

2.6 Cloning mutant alleles by gap-repair 33

2.7 Plasmid Recovery 34

2.8 Screening for mutants and overexpression screens. 36

Introduction to the cell cycle and cell growth. 37

3.1 Measuring cell number and volume

- The Coulter Counter 39

- The Sysmex- 40

- Haemocytometer 41

- Measuring cell size for individual cells. 41

3.2 Transition Point 42

3.3 Synchronising cultures with respect to the cell cycle

-Elutriation 43

-Arresting cells with ts-mutants: 44

-Arresting the cells with drugs 44

-Arresting cells in G1 by Nitrogen starvation. 45

-Arresting cells with pheromone 46

-Synchronous meiosis 46

3.4 Measuring DNA content of cells

- Fixing cells for flow cytometry 47

- Using the BD FACScan 48

- The diphenylamine reaction 50

4.1 Quick staining

- DAPI staining nuclei 53

- Staining the cell wall and septum with calcofluor 54

4.2 Indirect Immunofluorescence

- Guide 55

- Anti- tubulin immunofluorescence : formaldehyde 65

- Anti- tubulin immunofluorescence : methanol 67

- Actin rhodamine-phalloidin staining 68

- Actin immunofluorescence : methanol 69

4.3 ß-galactosidase detection 70

4.4 GFP 72

a. Nuclear isolation procedure 73

b. Preparing S. pombe chromosomal DNA 74

c. Chromatin extracts 77

d. Pulse Field Gel Electrophoresis 79

e. Genomic Digests 80

f. Southern blotting 80

g. Reprobing of Southern blots 82

5.2 Preparing probes by random priming 82

5.3 RNA preparation

4 ways to prepare S. pombe RNA 82

Northern blotting 84

5.4 Nuclear run on 85

5.5 Protein preparation

Large scale extracts 86

Small scale extracts 87

A. Native extracts 87

B. Denatured extracts 87

C. Boiled extracts 87

5.6 Immunoprecipitations 88

5.7 Histone kinase assays

A. Crude extracts 90

B. Immunoprecipitations 90

C. suc1p 90

5.8 In vivo protein labelling

35S-methionine labelling 91

32P-orthophosphate labelling 91

5.9 Bandshift assays with cell extracts 92

5.10 Antibody purification

Blot affinity purification of antibody 92

Affinity purification of anti-cdc13 antibodies fromthe SP4 crude serum 93

Purification of tea1 protein from E.coli 94

5.13 Expression of proteins in E.coli 99

Expression of his-tagged proteins 100

Expression of GST-tagged proteins, method I 101

Expression of GST-tagged proteins, method II 102

5.14 Transposon mutagenesis 104

You should put into the collection an aliquot of all plasmids you get from other labs or that you make whilst in this lab. To add a plasmid to the main collection all you need to do is give an aliquot to Nigel. If you need a particular plasmid you should take an aliquot from the collection and make your own prep. If a plasmid is missing or less than 50µl are left please let Nigel know.

Maps of all the plasmids are kept in a large black file on shelf in 402, when you add a plasmid to the collection please give a map and relevant information to Nigel.

We have several cDNA and genomic libraries. Information about them is in the plasmid file in 307 and aliquots can be obtained from Nigel. Any new libraries or ones obtained from outside should be added to the collection.

For polyclonal antibodies aliquots of sera are kept in the-70°C freezer in 3B. If you need affinity purified antibody you should prepare this yourself. Generally antibody is purified from 2-3mls of serum, this is then used by the lab until it is finished and then a fresh lot is prepared. This only works if everyone is prepared to take their turn doing the affinity purification. This also applies to the preparation of, for example, suc1 beads or HA beads. There are also aliquots of purified antibodies for most of the monoclonal antibodies.

Information about the antibodies is kept in a red file in 307 and on the computer in Lab Stuff- Cell cycle lab abs. If you make a new antibody whilst in the lab or get an antibody from elsewhere please put the information into this file and on to the computer and give the antibody to Nigel. All antibodies should at least be tested for western blotting and if possible for the ability to immunoprecipitate.

Some antibodies are in much shorter supply that others and it is best to check with Jacky before choosing one to work with.

We have an extensive collection of yeast strains that has been built up over many years and I cannot over emphasise the importance of adding any new mutants. It is essential that everyone adds new strains to the collection as they make them as well as making their own collection for when they leave. The collection is kept in the -70°C freezer in the lab and the backup collection is in the -70°C in 3B. Two glycerols should be made, one for the main and one for the backup collection. When you add a new strain to the collection the backup glycerol can be put into a box in the lab -70°C freezer, this is taken down to 3B every so often.

The strain should be entered in to the strain book AND into the

computer.

The strain collection is on the computer in Lab Stuff- Lab strains. Before you reisolate a strain from the collection please check the full details for any relevant information. If you need to add information about a strain you will need to use the password.

If you have any queries you should ask Jacky.

We have also some specific mutants that have not been put in the general collection. These include :

•Bea's bank - (screen for ts lethal mutants)

•Alison's mutants (uncharacterised from diploidising screens, e.g. septation mutants)

• Fulvia's weaker alleles. (weaker orb/tea mutants constructed by Fulvia)

• Val's mutants (shape mutants)

If you wish to use any of the above please speak to Chris and she will give you further assistance.

The lab has five general purpose Macintosh computers. Each member of the lab has a folder on the shared disk called Users in which you should keep your files these can be backed up on your optical disc. The lab also has two optical disk drives. Large files (such as image files) should be stored on optical disks and not in Users (except temporarily)

When you leave the lab you should get rid of your unwanted files and leave the remaining files in a folder called ex-members, in the lab optical disks. There are two copies of this folder each on a separate optical disk. If you have any problems you should ask Juan or Nigel.

There is an Endnote cell cycle / pombe database in Lab Stuff-lab master reference data. If any references that you need are not in this database you should add them when you use Endnote. In this way it keeps pretty much up to date. Please try not to duplicate entries, however there are already several duplicates but please don’t remove them if you find them. Some of the entries may contain some mistakes so if you find any please correct them.

The lab manual can be printed out from Lab Stuff- lab manual

A version is being prepared for the WEB.

Any books/manuals e.g. Maniatis etc. can be found in room 307 near the computer area. Feel free to borrow but please return after use and leave a note if you are going to be using for a long time. There is also a folder containing the product information that comes with kits and reagents. Again, if you use please put back after use and if you order something for your own use please add the product information to the folder. We also have a copy of "Current Protocols in Molecular Biology " on CD ROM. If you wish to utilise this - see Nigel.

Most media, solutions and sterile glassware are provided. If you require anything else you will need to make it yourself . There is no other technical help on a permanent basis. Chris will occasionally help with competent cells and plasmid preps but all requests for such help must go through Nigel.

Hopefully everyone will contribute to the smooth running of the lab. It is very helpful if everyone helps with amplifying libraries, making competent cells and ordering chemicals before they run out etc. Each person has a lab job and these change about every 6-12 months. Radioactivity is done once every two weeks by two people on a rotational basis. Everyone is on the rota even if they are not using radioactivity.

These are held each week, usually on Tuesday morning in Paul’s office. On average each person will do a lab meeting every 4-6 months. These meeting are informal and should not be regarded as formal seminars. Journal Clubs are held every 5th week. Before the lab meeting we have lab business which gives everyone an opportunity to pass on information, air their grievances etc. Because this is a big lab it is easy for an individual to miss announcements, so it is very important to attend the lab business so that everyone knows what is going on.

If you want to go to a national or international meeting you should fill in an application form which you can get from Liz. Paul will then decide whether you can apply to go to the meeting. Each ICRF lab has a limited budget for meetings/travel and everyone should try to get external funding. For graduate students the allowance it is approximately £1000 over 3-4 years. It is useful to join societies eg. The Genetical Society, Society for General Microbiology or Biochemical Society, as these are often a source of funding. All applications for money for meetings must go through Liz.

There are also regular day or evening meetings such as the Pombe Club, London Cell and Molecular Biology Club and DNA Replication Club. Information about these is displayed on notice boards around the building.

See Nigel who will explain everything you need to know.

ICRF has strict rules about taking equipment home and these should be adhered to for insurance purposes. We have two Macintosh power books that can be used at home for short periods. They should be brought back everytime you come into the lab and not left at your home. For more information see Liz.

A few months before you leave you should ensure that you have left all plasmids, libraries, strains, antibodies etc., that you have made, or requested whilst in the lab, with all the relevant information.

haploid cell: 12-14 µm at division

3.5µm width-constant

diploid 24µm length at division

5µm width-constant

average 2C DNA content for haploid wt 33.8fg/cell

average protein content for haploid wt 10pg/cell

average RNA content for haploid wt 3pg/cell (1-2% of which is polyA+)

1. Grow up cells in 0.8ml YES medium at 25-32°C for 2 days.

2. Mix with 0.8 ml of YES containing 50% glycerol (yellow freezing mix) in a cryotube. Place cultures at -70oC. The cells can then be stored at -70°C and remain viable for several years at least. It is wise to make a duplicate each time and store it in a different freezer. All new mutants or strains constructed should be deposited in the main laboratory collection.

When using cells requiring the active nmt promoter to survive, freeze them in supplemented minimal media and 50% glycerol.

For strains stored on glycerol at -70°C:

1. With a sterile spatula scrape off a small amount of frozen glycerol stock and patch onto a YES plate, keeping the stock frozen.

2. Incubate at 25°C-32°C for 1-4 days, depending on the strain.

3. When growth is visible streak out to single colonies on YES and incubate at 25°C-32°C for 2-3 days.

Strains stored as slants or patches are streaked out onto YES plates directly, and incubated at 25°C-32°C as appropriate.

It is essential to check the phenotype of any re-isolated strain before carrying out any genetic or molecular procedures.

1. ploidy

2. mating type

3. testing the genotype by PCR

4. temperature/ cold sensitivity

5. auxotrophy

6. adenine mutants

It is important to check the ploidy because certain strains of S. pombe diploidise at a high frequency. Haploid cells divide at approximately 12-15 µm in length and are 3-4 µm in width. Diploid cells are both longer (20-25 µm at division) and wider (4-5µm); they are also less viable than haploid cells and a diploid colony contains more dead cells (1-5%). Phloxin B is a stain that accumulates in dead cells, which become stained dark red. By growing a strain in YEP plates it is possible to screen for haploid colonies, which will be stained light pink whereas diploid colonies will be darker pink. This can be confirmed by microscopic examination of the cells. Streaking of strains on a plate next to a known haploid can help identification by having cells of both types in the same microscopic field.

To check mating type the strain should be crossed to h+ and h- tester strains (see later) and tested as above. PCR can also be used see below.

Also see following protocol (Determination of mating type by PCR) for a an alternative protocol

PCR mix: prepare enough for the number of PCR reactions planned + 1

per reaction: 42 µl water

(Note: 10x Vent buffer from NEB may be used instead)

0.5 µl 10 mM each dNTP

1 µl 50 µM primer 1

1 µl 50 µM primer 2

49.5 µl

1. pick colony and resuspend well in 30 µl PCR mix in a 0.5 ml PCR tube. Total amount of yeast is a "match head full" and solution will be cloudy upon resuspension.

2. boil 5 min (use PCR machine: 5 min 100 °C, then 4 °C)

5. amplify 30 cycles: 94 °C 30 sec / 50 °C 30 sec / 72 °C 30 sec

Note:

a) optimal annealing temperature may vary depending on Tm of primers

b) allow approximately 1 min at 72 °C per kb of extension.

6. extend final cycle at 72°C for 10 min to ensure complete extension of products, then 4 °C.

7. remove 10 - 15 µl and check on a gel. Gel purified band can be directly sequenced.

(This protocol doesn't work with Vent polymerase)

Using mat1-P and mat1-M specific primers, it is possible to determine the mating type of a strain even if it is sterile. With common primer MT1, the mat1-P specific primer MP or mat1-M specific primer MM produces a characteristic PCR product, 987bp or 729bp respectively. h+/h- diploid cells or h90 cells give both bands. The sequence of the primers are MT1:AGAAGAGAGAGTAGTTGAAG , MP:ACGGTAGTCATCGGTCTTCC, and MM:TACGTTCAGTAGACGTAGTG.

Use 20ul for each tube and add 1ul of template DNA prepared from pombe colony or culture. Instead of preparing DNA, picked fresh colony can be used if it is denatured in a small amount (~5ul) of water at 95C for 10min. Add 10ul mineral oil on top of each reaction.

Reaction mixture

H2O........................76ul

X10 buffer..............10ul

2mM dNTPs..........10ul

50uMMT1primer....1ul

50uMMPprimer......1ul

50uMMMprimer.....1ul

Ampli Taq Pol.........1ul

--------------------------------

total ..........100ul

PCR cycle

step1 94C 4min

step2 94C 30sec

step3 52C 30sec

step4 72C 2min

repeat step2~step4 for 30cycles

step5 72C 5min

After the reaction, add 2ul of loading buffer and ~20ul of chloroform. Analyze in an agarose gel.

4) Temperature/cold sensitivity.

Many temperature sensitive (ts) and cold sensitive (cs) mutants have been isolated in S. pombe. They can be checked by replica plating onto YEP (YES + phloxin B) and incubating at the restrictive temperature. Phloxin B will stain the dead cells and these can be examined under the light microscope for checking the ts /cs phenotype.

The auxotrophic markers most commonly used in S. pombe require adenine, glutamic acid, histidine, leucine, lysine and uracil, although others are available. We use 225mg/L (12ml of 7.5mg/ml stock solution per 400ml medium, or, for uracil, 24ml of a 3.75mg/ml stock per 400ml medium) although it is OK to go down to 75mg/L except when growing leu- strains, which are slow to get going under these conditions. To test for auxotrophy the strain is grown up to single colonies on YES and then replica plated to minimal medium with and without the appropriate supplement. The plates are incubated for 1-2 days and then examined for growth under these conditions.

Strains marked with auxotrophic markers are often compromised for growth even in the presence of the appropriate supplement.. e.g. leu- cells are slow to exit from starvation and ura- cells have some cell wall defects. For this reason it is best to cross out all unnecessary markers when carrying out physiological experiments.

Certain alleles of ade6 turn pink when grown on low adenine medium (10mg/L, or 0.5ml of a 7.5mg/ml stock solution per 400ml medium), due to the accumulation of a metabolic precursor. Ade6-M210 turns darker than ade6-M216, ade6-704 is the darkest of all.

Haploid strains of S. pombe grow with the following generation times:

(Diploid strains grow similarly)

YE 25 3h

29 2h 30min

32 2h 10min

35.5 2h

minimal 25 4h

29 3h

32 2h 30min

35.5 2h 20min

T = log (2t2-t1) T = generation time

---------- y = cells/ml at time t2

log (y/x) x = cells/ml at time t1

The relationship between OD and cell number is linear up to about OD 1.

To generate cultures in mid-exponential growth use a fresh colony or overnight patch of a strain of checked phenotype to inoculate 10 ml YES (or minimal medium if the strain carries a plasmid with selectable marker) and incubate for 1-2 days at the appropriate temperature until cells are in early stationary phase. ( See note at the end of the page)This pre-culture can then be used to inoculate a larger culture, taking into consideration the generation times .

The following formula can be used to calculate the volume of preculture from which to generate a larger, overnight culture, which is grown up with shaking:

_________________

OD

Flask size should be selected according to the required volume of culture: 25ml flasks for up to 10ml culture, 100ml flasks for up to 50ml, 200ml for 100ml cultures, 250ml for 125ml cultures and 500ml for 250ml cultures. It is important not to inoculate stationary cultures at an O.D. of less than about 0.05, because S. pombe cells grow better together or in conditioned media and are often reluctant to get going when alone in liquid culture.

The media for growing S. pombe strains are given in Table I. YES liquid or agar containing solid medium is used for vegetative growth. Yeast extract from most sources when used in YES inhibits conjugation and sporulation. EMM2 minimal media is synthetic and therefore highly reproducible. For this reason it is recommended for physiological experiments with the required supplements added. YES is however prefered for certain application: for actin staining, branching experiment and certain blocks (eg nda3).

Note: Certain strains have difficulty in exiting from stationary phase (cdc25, ura-)therefore it is best to inoculate from mid-exponential phase. Not shaking the cells in preculture could also help because it stops cells entering late stationary phase.

When inoculating into EMM from mid exponential YE culture, it might be better to spin the cells from the preculture down, in order to reduce the amount of YE carried over.

Conjugation and sporulation cannot take place in S. pombe except under conditions of nutrient starvation. EMM lacking nitrogen and containing 1g/l sodium glutamate is generally used for genetic crosses. See note for alternative media.

Zygotic ascus Azygotic ascus

A cross between a homothallic and a heterothallic strain mostly generates asci of the homothallic parent. This is because yeast cells prefer to mate with sister cells which are of different mating type due to homothallic switching. Mixing cells in a 10:1 ratio of the heterothallic:homothallic strain will increase the likelyhood of a cross between the two strains and is usually sufficient for strain construction.

If conjugation is very poor then it is preferable to first select a diploid hybrid between these strains (see below) and then subject this to tetrad or random spore analysis.

Asci are placed in a line about 3mm apart on a YES plate using a micromanipulator (we have two: one from Singer-MSM which uses an inverted plate and is automated and one from Zeiss Jena which uses upright plates and spores are dragged across the plate rather than lifter off the plate). The asci walls are then left to breakdown at 37°C for about 3-5 hours, or at 20°C overnight. It is usually a good idea to isolate more asci than you will need to dissect at the end, as some may not breakdown their cell wall or do so very slowly. Each ascus is then micromanipulated to give a line of four isolated spores, separated by about 3-5mm. The spores are incubated until colonies form at the appropriate temperature for the cross. Digestion of the ascus wall with enzymatic treatment is not necessary since the spores easily fall apart giving free spores. Also, unlike S. cerevisiae spores, S. pombe spores do not stick to each other. This means the technique of tetrad dissection is generally easier than with S. cerevisiae.

Using a three day old cross check for the presence of asci under the light microscope. Random spore analysis allows many more spores to be examined than in tetrad analysis and in this way recombination mapping and strain construction can be carried out. However, it is important that all the classes of spores are viable when studying recombination frequencies, otherwise tetrad analysis becomes necessary.

1 ml of sterile distilled water is inoculated with a loopful of the cross, 20ul of a 1 in 10 dilution of Helicase (Helix pomatia juice) is added and the mixture incubated overnight at 25-29°C or for at least 6 hours at 29°C. Helicase is a crude snail gut enzyme that breaks down the ascus wall and kills vegetative cells. 5ul of glusulase in 1 ml of sterile distilled water can be substituted for the Helicase.

The spore number/ml is counted using a haemocytometer. Between 200-1000 spores/plate can be plated out on YES Agar or selective medium. The plates are then incubated until colonies form.

NB: if the cross works reasonably well 100uL of a 1:1000 dilution will give approx. 200 colonies per plate if you inoculate the 1ml culture with a generous loopful of cross. Easier than counting, especially if you are doing large numbers of crosses.

A diploid is constructed (see below) from 2 recessive mutations. If the diploid is wild type, genetic complementation is taking place and it can usually be assumed that the two mutations are in different genes. It should be borne in mind when doing these types of studies that it is possible to get intragenic recombination in large genes where several protein subunits are involved.

These can usually be made by free spore analysis and selecting microscopically for the required phenotype (or by replica plating where suitable nutritional markers are involved). Where the double mutant cannot be distinguished so easily (or where it is not known what the phenotype of the double mutant will be),tetrad analysis is carried out on the meiotic progeny of a cross between the two strains of interest. Three types of tetrad are normally produced: the parental ditype (PD) if there is no recombination, the tetratype (TT) if there is single crossing -over (i.e. two recombinants are produced per ascus) and the non-parental ditype (NPD) if there is double crossing-over (ie. all recombinants are produced), or if the two genes are on different chromosomes, (by random assortment on the metaphase plate).

It is usually possible to determine the types of tetrads produced, in say a cross between two ts strains, by examining the phenotype after replica plating to phloxin B at the restrictive temperature. Where non-parental ditypes are produced, two spores will normally be wild-type, and two will be the double mutant, and thus the double mutant can easily be identified in this type of tetrad. In parental ditypes, none of the progeny will be double mutants, and in tetratypes 1 out of the 4 will be the double mutant (3 will be ts assuming the double mutant is ts) and 1 will be wild-type). It will sometimes be possible to determine which is the double mutant by microscopic examination at the restrictive temperature. Tetrad types are normally produced in the proportions 1PD: 1 NPD: 4TT for markers that are unlinked but on the same chromosome. If they are on different chromosomes the number of tetratypes depends on the distance of the loci from the centromere. A predominance of parental ditypes suggests linkage.. The presence of a double mutant can be confirmed by back-crossing if the single mutants are phenotypically different. A double mutant backcross will give a 3:1 segregation of mutant:WT. If a single mutant is backcrossed, there will be a 2:2 segregation.

This is achieved by the construction of diploids containing both a mutant and a wild type copy of the gene to be tested; if the mutant phenotype is not expressed in the diploid , then it is usually assumed that the mutation is recessive, whilst expression suggests the presence of a dominant mutation; however, unless very large numbers of diploids are tested in this way, it is wise to check that growth/lack of growth is not due to the presence of a homozygous diploid possessing 2/0 copies of a wild type/ mutant gene - such events occur in S.pombe at low frequency by recombination between replicated chromosomes in the zygote. The frequency with which such events occur is affected by map position - recombination frequency decreases with increasing distance from the centromere, and also at certain recombination "hotspots", for example near the mating type locus on chromosome II. This could easily be sorted out by pulling tetrads on the diploid under study. In addition, the diploid may be subjected to random spore analysis to check that 50% of the spores have the wild type phenotype and the remaining 50% have the mutant phenotype.

Another important use of this approach is if 2 mutants are very closely linked and you want to know whether or not they are in the same gene. A diploid is made from the 2 strains and complementation tested under restrictive conditions. Growth indicates close linkage but different genes. Again, tetrads on such a diploid would confirm that the genotype is as predicted.

Diploid cells arise spontaneously in most S. pombe strains probably as a result of endoreplication. This characteristic can be used to isolate homozygous diploids of any strain. The strain is streaked out to single colonies on YEP (YE + Phloxin B) agar and incubated until colonies form. Diploids can be identified as clones that stain dark red with Phloxin B and which contain large cells upon microscopic examination. These diploids can undergo a diploid mitotic cycle and when starved of nutrients can conjugate with diploid cells of the opposite mating type to form a tetraploid zygote. This can be sporulated to generate four diploid spores, although aberrant segregation of chromosomes can occur during the tetraploid meiosis.

Homothallic strains also generate diploid cells which can switch mating type and therefore be induced to undergo meiosis. The cells can directly produce azygotic asci without mating and thus can be unambiguously identified. (They can also mate with each other)

Sporulating diploids can be isolated by crossing h- and h+ haploids with complementary growth requirements, for example using strains with the markers leu1-32 h- and a ura4-d18 h+. At various time-points after crossing (e.g. 8, 12, 24 hours), when conjugation has occurred, a loopful of the cross is streaked out onto minimal medium which will only allow growth of a conjugated diploid. However, since mating is rapidly followed by meiosis and sporulation, many of the colonies growing on selective media will be prototrophic haploid segregants. To circumvent this problem, ade6-M210 and ade6-M216 mutations are commonly used. ade6-M216 colonies are light pink and ade6-M210 dark pink on plates of YE medium or EMM containing 10mg/L adenine because of accumulation of a red adenine precursor. On media containing adenine in excess of 10mg/L the red colour is not observed. Diploid cells containing both mutations grow in the absence of adenine and the colonies formed are white due to intragenic complementation between the two ade6 alleles. Because the alleles are tightly linked and there is infrequent gene conversion, spores generated by meiosis are unlikely to be adenine prototrophs and will not form colonies on the selective media.

Sometimes it is required to cross a diploid and a haploid strain. Such a cross yields less than 10% spore viability and most of the segregants are slow growing because of aneuploidy. Since S. pombe has only three linkage groups, normal haploid and diploid segregants can be obtained at a reasonable frequency from random spores; they are easily identified since they grow well.

Protocol for generating an h+/h- sporulating diploid

4) There will usually be two types of colonies on the YE plate: larger, white colonies (these are diploids) and smaller, red colonies (these are haploid colonies ade-which grew from spores generated when the diploid reached stationary phase on the EMM -ade plate and underwent sporulation, they are red because YE contains low levels of adenine and ade- cells will grow but accumulate a red metabolic precursor ). Check under the microscope to make sure that cells in the darker colonies have a larger size. Patch the diploids on YES and replica plate the YE plate on EMM-glutamate. Check after 2-3 days that the diploids picked are sporulating.

Remember to propagate the diploid on YES, which inhibites sporulation.

If the diploid is to be used for deletion or tagging of a gene it is a good idea to dissect a couple of tetrads beforehand to make sure that no recessive mutation is present in the diploid.

This technique is required if crosses between two strains will not occur and a diploid is desired.

0.65 M Kcl

1M sorbitol

30% PEG6000

Novozyme

Grow cells to mid-log phase in minimal medium (nb MIN with low glucose is supposed to make the cell wall weaker but in my experience it is not necessary and some strains don't like it)

Harvest cells, wash twice with water and once with 0.65M KCl.

Resuspend pellet in 0.65M KCl containing 1 mg/ml Novozyme, at about 108 cell/ml.

Incubate at 30oC.

Check after about 15 min, you should see spheroplasts. If not, you can increase the enzyme concentration, but it's advisable to start slowly with unknown strains.

Continue incubation in Novozyme until protoplasts are formed. It's not enough to have spheroplasts!

Wash protoplasts three times with 1M sorbitol.

Note: spin them at 2000 RPM, 8 min. Do NOT vortex to resuspend, it can be done with a loop in a small volume first and then by gently shaking the tube in a larger volume.

Note: mix it just before use, do NOT autoclave together, because it precipitates.

Incubate at room temperature for 20min. Plate onto selective medium containing 1M sorbitol.

Note: plating in top agar (same as selective, with 1% agar) increases regeneration frequency, although makes it impossible to replicaplate colonies.

1. Grow cells in MML (0.5% Glucose + thiamine) to late log-phase. Since total 1-5 * 107 cells are required, more than 10 ml should be cultured.

2. Measure OD at 595 nm.

4. Mix cells with equal cell number and total 1ml.

5. Add 0.2 ml Novozyme234soln (bufferA + 5 mg/ml Novozyme)

You can increase Novozyme concentration to 5 mg/ml as final.

6. Incubate 25-30˙C for 1hr with rotating. After incubation, you should check whether Novozyme works well under microscopy and confirm that most of the cells are round after addition of equal amount of H2O. Usually 20-70% of the cells were round.

7. Spin down at 2000rpm for 15 sec. If the pellet is not visible or soft, change the angle of the tube and spin down again.

8. Discard the sup and add 1 ml of bufferB (1.2 M sorbitol, 30mM Tris-HCl pH7.6) to the tube.

9. Spin sown again and discard the sup and add 1 ml of bufferB.

10. Spin sown again and discard the sup and add 0.3 ml PEG buffer (30% PEG4000, 10 mM Tris-HCl pH7.6, 10mM CaCl2). Be careful not to suspend the pellet.

11. Incubate at 25-30 ˙C with standing for 30 min.

12. Discard the 0.15ml of sup and plate the cells on SD sorbitol plate (0.67% SD, 1% glucose) and incubate 25-30˙C for 5-14 days.

1.2 M sorbitol (H2O 830 ml + sorbitol 220g)

Why bother putting your gene on the genetic map? There are several reasons.

1) First of all if a gene cannot be isolated by classical complementation of a mutant phenotype, it is possible to clone it by physical mapping. Initially by crude mapping and then by the use of ordered cosmids to rescue the function and to locate the gene. The mutated gene responsible for a subtle phenotype can be isolated in this way.

2) In addition, a mutation with an interesting phenotype may have already been identified that lies in the same locus as your gene and which sheads light on its function.

3) Mapping completes the genetic characterisation of your gene. However, there is admittedly limited information gained, and most people now seem to map their gene by accident, because it shows linkage to some other locus present when strain constructions are carried out.

The first step to mapping your gene is to assign it to a chromosome. There are several ways to do this. The older way requires chromosome loss expts. You construct a non-sporulating diploid with differently marked chromosomes. You treat it with fluorophenylalanine (FPA) , a drug which promotes chromosome loss. If using minimal, use 0.04% of m-FPA, and if using YEA, use 0.1% p-FPA. Some workers consider that m-FPA is less toxic, and therefore the agent of choice. Because pombe rapidly haploidises when one chromosome is lost, you end up associating your gene with the presence or absence of one of those marked chromosomes.

Another option is to use a swi5 strain. A swi5 mutant is not only defective in switching (hence the name) but has a general reduction in recombination frequency. That is, with a five fold reduction in recombination frequency, distant linkage that would otherwise be invisible can be seen. Thus with just a few markers in the background, the gene of interest is likely to show linkage to at least one. Practically, this means crossing the gene of interest into a swi5 background, and mapping against a swi5 strain with markers on all chromosomes.

The most straightforward way of assigning a cloned gene to a chromosome is getting a strip of a chromosome blot and probing it by standard Southern methods. Since pombe has only 3 chromosomes, this gives an unambiguous result.

Once you have a chromosome assignment, you have to map your gene against the markers on that chromosome. The swi5 technique can be very useful here, as you can put a variety of markers on, say, chromosome II into the cross, and rapidly localise the gene of interest. Or, you can cross against a multiply marked strain and look for linkage that way. Practically speaking, it is hard to see linkage farther than about 40-50cM in a wildtype (eg, non-swi5) background. Most mapping functions are based on tetrads. This is because the non-parental ditype class gives a powerful estimate of the frequency of multiple crossovers, which are otherwise silent. When analysing tetrad data, a cross of only 10 tetrads usually gives enough information to indicate linkage. If markers in a cross A B x a b can assort randomly, then the following classes are expected:

Parental Ditype (PD) Nonparental ditype (NPD) Tetratype (T)

A B A b Ab

A B A b a B

a b a B A B

a b a B a b

The NPD class involves at least two crossovers. For the number of PD to equal to the number of NPD, the two genes must be unlinked. For completely unlinked genes, neither of which is linked to its centromere, the expected frequency is 1PD: 1NPD: 4T.

Therefore, even with normal statistical variation, if you see a high frequency of T and some NPD in your 10 tetrads, the two genes are probably unlinked. If you start to see a bias towards the PD class, it is worth pulling a few more.

If your gene is linked to a centromere, it falls into a special class. Centromere linkage can be determined by one cross, even if you don't know what chromosome the gene is on. The theory is as follows: if a gene (eg, lys2) is tightly linked to its centromere, then there is little chance of a crossover between that gene and centromere. This means that the chromosome with that marker will always segregate from its homologue in meiosis I. Therefore, if you cross TWO centromere linked genes, they can only give you ditype progeny, and no tetratypes, because the formation of tetratypes requires a crossover between marker and centromere that allows meiosis II segregation. Expanding this further, if you cross your gene to a tightly-centromere-linked marker, you can calculate the distance of your gene from its OWN centromere, even without knowing which chromosome your gene is on. lys2 is tightly centromere linked on chromosome I.

S. pombe does not suffer interference. This means that, unlike the case in budding yeast, the fact a crossover has occurred doesn't seem to affect the likelihood of additional crossovers in the same interval. There are a few exceptions to this rule with some recombinational hotspots, but basically it means that the usual mapping functions give a reasonable approximation of distance. The standard formula is the Perkins formula:

(T + 6NPD)

distance in cM = ----------------------

(PD + NPD + T)

This formula tends to under estimate the actual distance, because of multiple crossovers. In practise, anything greater than 60cM is unreliable. It is useful to compare the Perkins answer with the Papazian answer, which uses the Poisson distribution to estimate the number of multiple crossovers:

(PD - NPD)

distance in cM = 100 X -0.5 ln ----------------------

(PD + NPD + T)

The answers should be fairly comparable.

Now that cosmid and P1 libraries are available physical mapping is also possible.

Both EMS and nitrosoguanidine can be used for mutagenesis, the former is safer to use but nitrosoguanidine is a more effective mutagen. Nitrosoguanidine tends to be the agent of choice.

1) First do a survival curve 50% survival is often used for

mutagenesis as there is approximately one hit per gene

and wash 1X in TM and 2X in saline. You may need to spin the cells from 2-4min to ensure a good pellet.

100ml of each on min medium and incubate at the permissive temperature until colonies appear.

The percentage of survivors is usually approximately 40%, 15% and 3% for 30, 60 and 90 minutes, respectively (these percentages are strain dependent). A survival curve can be plotted by plating 100 and 1000 cells/plate and plotting % survivors as a function of exposure time to the mutagen. The length of exposure required to produce the desired level of lethality can the be determined and used in subsequent experiments

8) Using the appropriate time point repeat the mutagenesis using

9) To save repeating the mutagenesis you can take sufficient cells at what you think will be the right time point(s) and keep them in 1ml EMM2 at +4°C until you have done the survival curve. Cells lose viability but keep for a week or so. I do the survival curve after leaving the cells in the cold room overnight.

10) After mutagenesis it is advisable to backcross the strains at least three times to check for the presence of double mutants etc., especially if the mutagenesis has been heavy (>40% lethality).

11) NG is inactivated by soaking any contaminated materials in 0.1M HCl overnight.

ALL MANIPULATIONS WITH NITROSOGUANIDINE (INCLUDING INACTIVATION) MUST BE CARRIED OUT IN A FUME HOOD !!

Edinburgh Minimal Medium (EMM 2) (Used for vegetative growth)

3 g/l potassium hydrogen phthallate (14.7mM)

2% (w/v) glucose (111 mM)

20 ml/l salts (stock x 50)

1 ml/l vitamins (stock x 1000)

0.1 ml/l minerals (stock x 10,000)

Salts x 50

50 g/l KCl (0.67 M)

Vitamins x 1000

1 g/l pantothenic acid (4.20 mM)

10 g/l nicotinic acid (8l.2 mM)

10 g/l inositol (55.5 mM)

10 mg/l biotin (40.8 µM)

Minerals x 10,000

5 g/l boric acid (80.9 mM)

0.4 g/l molybdic acid (2.47 mM)

1 g/l KI (6.02 mM)

10 g/l citric acid (47.6 mM)

After autoclaving, a few drops

of preservative is added. (1:1:2, chlorobenzene : dichloroethane :chlorobutane)

Minimal supplemented (Used for vegetative growth)

EMM2 + 225 mg/l supplements as required.

Minimal low glucose (Used for Yeast transformations)

As EMM but 0.5% (w/v) glucose instead of 2% (w/v) .

Minimal sorbitol (EMMS - Used to grow up transformants)

As EMM. Add 1.2M sorbitol.

Minimal glutamate (EMMG Used for sporulating diploids in liquid).

0.5% (w/v) Oxoid yeast extract.

3.0% (w/v) glucose.

Yeast extract + supplements (YES - Vegetative growth)

YE+225 mg/l adenine, histidine, leucine, uracil and lysine hydrochloride.

Yeast extract + phloxin B (YEP - Checking ploidy)

YES+5 mg/l phloxin B (Sigma No. P 4030). Add when the medium has cooled below 60°C from a 5g/l stock solution in sterile distilled water.

3% (w/v) Bacto-malt extract (DiaMalt AG or Difco). Supplements added as for YES except lysine. Adjust to pH 5.5 with NaOH.

Solid media is made by adding 2% Difco Bacto Agar.

All media is prepared in bulk. It is sterilized by autoclaving at 10 psi for 20 min. At this pressure very little caramelization of glucose takes place. The media is stored in 500 ml bottles and agar is remelted in a microwave oven before using.

S. pombe plasmids consist of a bacterial origin of replication and selectable marker, a yeast selectable marker and an equivalent to an autonomous replication sequence (ars) which is responsible for high frequency of transformation.

Budding yeast markers used in S. pombe are the LEU2 and URA3 genes. Plasmids containing these markers complement the S. pombe mutations leu1- and ura 4-. The URA3 gene is expressed very poorly in S. pombe and does not rescue the ura 4- mutation when it is present as a single copy or even at moderate levels. S. pombe markers commonly used are ura 4+, sup3-5., leu1, his3, and his7.

In contrast to S.cerevisiae, in S. pombe a bacterial plasmid such as pBR322 carrying a marker gene such as LEU2 is able to replicate often to high copy number. However, the transformation frequency obtained when using such plasmids is very low. The addition of S. pombe ars1+ sequences or the S.cerevisiae 2 µm origin leads to high frequency of transformation and reduction in the copy number. So it seems that in S. pombe high frequency of transformation and effective replication capacity are to some extent independent phenomena.

Plasmid vectors based on 2 µm (pDB248, YEp13) are mitotically unstable, their copy number is low, they are much more prone to rearrangements (tandem duplications or deletions) and they are more difficult to recover from fission yeast than plasmids carrying S. pombe ars1+. Plasmids containing ars1+ are also very unstable (with the exception of pFL20 and pMB332); their copy number is higher and they tend to produce polymers with various numbers of repeats units. pFL20 and pMB332 yield rather stable transformants both mitotically and meiotically, due to the presence of a stb (stable) element . This element is not an ars sequence nor it is a centromeric sequence. Plasmids containing this element still segregated asymmetrically ten times more frequently during mitosis than S.cerevisiae CEN plasmids.

Plasmids derived from the ones described above have been used to increase the expression of certain gene products. pSM1 and pSM2 are derivates from pDB248, made by inserting the SV40 early promoter . Genes linked to this promoter are expressed at moderate levels. pEVP11 contains the S. pombe adh+ promoter inserted into YEp13 S.cerevisiae vector. pART1 and pMB332 also have the S. pombe adh+ promoter inserted into pIRT2 and pFL20, respectively. The adh1 promoter is very active - about 5-20 x greater than the SV40 early promoter. pIRT2 is based on pUC118 and has S.pombe ars1 and S.cerevisiae LEU2 inserted.

Recently, plasmids containing inducible promoters have been developed. These include various plasmids containing the thiamine repressible promoter, as developed by Maundrell (1990). Several versions are available in which the promoter sequences have been mutated to different degrees to give lower levels of expression (see Forsburg ref). pREP1 contains the wild type promoter. There is a significant background expression level ie with thiamine. The induced level is about 80X greater than the repressed level and about 6X greater than the level produced by the adh1 promoter. pREP41 has a 6X lower induced level and a 15X lower repressed level than the wildtype promoter. pREP81 has an induced level about 80X lower than the wildtype promoter (comparable to the repressed level of the wildtype promoter), whilst in the absence of thiamine the level is reduced a further 250 fold.

Reference Hoffman & Winston 1989, Gene 84 473-479

This is a URA3 (S. cerevisiae) based vector pCHY21. It has the glucose repressible promoter of the fbp (fructose bisphosphase) gene upstream of a polylinker with 4 unique sites, Xho1, BamH1, PvuII and Nhe1. The promoter is OFF in 8% glucose and ON in 0.1% glucose +3% maltose.

The vector has been tested with the cdc13∆90 gene and worked well. All the cells became blocked in mitosis with a cut phenotype by 5 hours after shifting to 0.1% glucose 3% maltose

The average induction of fbp::lacZ is

repressed derepressed

30 +/- 3 4405 +/-417

Intermediate expression can be obtained using

b gal expression

3.0% maltose 523 +/-78

0.2% glucose 289 +/- 37

0.5% glucose 88 +/- 12

1% glucose 55 +/- 6

1) Promoter is derepressed in stationary phase

2) URA3 is not well expressed when integrated in S. pombe

3 Wild type cells do not grow well in 0.1% glucose+3% maltose

1) Grow cells to an appropriate OD in minimal medium +8% glucose.

2) Wash the cells X2 with 0.1% glucose + 3% maltose minimal medium.

3) Resuspend the cells in 0.1% glucose + 3% maltose minimal medium and follow induction. The promoter is derepressed 30 fold by about 5h.

4) The cells should not be allowed to enter stationary phase

2. Harvest the cells at 3000 rpm for 5 minutes at room temperature.

3. Wash cells in 40 ml of H2O and spin them down as before.

acid) and dispense 100 µl aliquots into Eppendorf tubes. Incubate at 30°C (25°C for ts mutants) for 60 - 120 minutes. Cells will sediment at this stage.

5. Add 1 µg of plasmid DNA in 15 µl TE (pH 7.5) to each tube and mix by gentle vortexing, completely resuspending cells sedimented during the incubation. Do not allow the tubes to cool down at this stage. Add 290 µl of 50 % (w/v) PEG 4000 prewarmed at 30°C (25°C for ts mutants). Mix by gentle vortexing and incubate at 30°C (25°C for ts mutants) for 60 minutes.

6. Heat shock at 43°C for 15 minutes. Cool the tubes to room temperature for 10 minutes.

7. Centrifuge at 5000 rpm for 2 minutes in an Eppendorf centrifuge. Carefully remove the supernatant by aspiration.

8. Resuspend the cells in 1 ml of 1/2 YE5S by pipetting up and down with a pipetman P1000.

9. Transfer the suspension to a 50 ml flask and dilute with 9 ml of 1/2 YE5S. Incubate with shaking at 32 °C (25°C for ts mutants) for 60 minutes or longer.

11. Plate aliquots of less than 0.3 ml onto minimal plates. If necessary, centrifuge the cells at this stage and resuspend in 1ml of media to spread more cells on a plate.

Some strains may grow poorly or not at all in MB medium; URA 4 strains in combination with a temperature sensitive CDC allele seem to be particularly intractable. It is possible in some cases to overcome this by the following procedure:

1. From a preculture in YE5S inoculate a 25ml culture of YE5S and grow until in early exponential phase.

2. Harvest cells and resuspend in 200ml minimal medium (with appropriate supplements); grow again till early exponential phase.

3. Harvest cells and resuspend in MB medium for the last 1-2 cell divisions.

1. Grow fission yeast cells in MM to 1x107 cells/ml.

2. Pellet 50ml of cells per transformation.

3. Wash cells in 50ml sterile water. Transfer to eppis in 1ml water. Wash in 1ml of LiAc-TE.

4. Resuspend in LiAc-TE at 2x109 cells/ml (1/200 original volume).

5. Mix 100µl cells with 2µl carrier DNA at 10mg/ml and up to 10µl of DNA; mix gently.

6. Incubate at RT for 10 min.

7. Add 260µl of 40% PEG/LiAc-TE; mix gently.

8. Incubate 30-60 min at 29°C-30°C, or lower for temp. sensitive strains.

9. Add 43µl pre-warmed DMSO; mix gently.

10. Heat shock at 42°C for 5 min.

11. Pellet and wash once with 1ml water.

12. Pellet and resuspend in 500µl water and plate 250µl in duplicate.

LiAc-TE: 0.1M lithium acetate, 10mM Tris pH 7.5, 1 mM EDTA

Carrier DNA: boiled sperm DNA 10mg/ml

LiAc-TE-PEG: LiAc-TE plus 40% PEG4000

4. 40-100 µl of the cell suspension are added to chilled cuvettes containing the DNA for transformation ( 100 ng ) and incubated on ice for 5 minutes.

5. Dry bottom of cuvette and pulse cells. The electroporator (Jensen) is set to 1.5 kV, 132 Ω, 40 µF. For a BioRad electroporator the settings are 1.5kV, 200 Ω, 25µF.

6. Cells and DNA are transferred to a pre-chilled cuvette and pulsed; 0.9 ml of ice-cold 1M sorbitol is then immediately added to the cuvette; the cell suspension is then returned to the Eppendorf and placed on ice while other electroporations are carried out.

Note: This method does not seem to work very well with the sup 3-5 marker.

2. Harvest cells (at 2000rpm, 5 minutes, J-6), decant supernatant and resuspend the pellet in 10 ml of:

40mM EDTA pH 8.0

transfer to 50 ml round bottom tube, e.g. Oakridge tubes.

3. Harvest cells and resuspend each tube in 5 ml of:

1.2M Sorbitol. Adjust to pH 5.6 with 5M NaOH.

and add 25 mg NovoZymTM 234 (added after autoclaving). Incubate at 37°C for 15-30 minutes (check under the microscope) until spheroplasts have formed.

4. Add 35 ml of:

10mM Tris-HCl pH 7.6

1.2M sorbitol

5. Wash twice more in 20 ml each time resuspending gently in 1ml first.

At the last resuspension take a sample and count the number of protoplasts with a haemocytometer.

10mM Tris HCl pH 7.6

1.2M Sorbitol

and combine the tubes.

7. Using 100 µl protoplast/ transformation add 1-10 µg of transforming plasmid in up to 1/10 total volume. Incubate at room temperature for up to 60 minutes.

8. Add 1ml of:

10mM Tris-HCl pH 7.6

20% PEG 4000

and incubate at room temperature for 15 minutes.

9. Spin at 2000 rpm for 5 minutes, drain well and resuspend the protoplast in 0.2-0.5 ml of:

10mM Tris-HCl pH 7.6

1.2M sorbitol

0.5mg/ml Yeast extract

5µg/ml supplements ( leu, ura, ade, his)

Incubate at 30°C for 30-60 minutes.

2. Use 10 ml of this culture to inoculate 40 ml of fresh YEPD medium and incubate for 4-5 hours.

3. Harvest cells at 3000 rpm for 5 minutes. Wash once in sterile distilled water and resuspend in 0.6 ml of buffer I (20 mM Tris-HCl pH 7.5, 2 mM EDTA, 0.2 M LiCl) to give a total volume of about 1.2 ml and a final concentration of 2 x 109 cells/ml.

4. Incubate at 30°C for 1 hour with gentle shaking.

6. Add 700 µl of buffer II (40% PEG 4000, 0.1M Li-Cl in TE buffer, sterilised with 0.2 µm filters). Mix by inverting the tube gently. Incubate at 30°C for 30 minutes.

7. Heat shock at 46°C for 25 minutes.

8. Spread the cell suspension directly into YNB (0.67% yeast nitrogen base without aminoacids, 2% glucose, 1,5% agar). Colonies appear after four to six days at 30°C.

In S. pombe integration by homologous recombination is usually more frequent than non-homologous recombination; but for certain loci homologous recombination may only represent about 5-10% of the integration events. On average about 0.1% of the transformants obtained after transformation with an ars plasmid will have an integrated copy of the plasmid at the homologous locus. The frequency of integration can be enhanced up to ten fold by a single cut of the plasmid in the region of interest to facilitate the recombination event.

There can be problems concerning the selective markers used that may complicate the integration of a plasmid in S. pombe. The S.cerevisiae URA3 gene on a multicopy plasmid complements S. pombe ura 4- mutations, but is poorly expressed in S. pombe and most of the integrated versions of URA3 fail to complement ura 4- mutations. Therefore LEU2 (does not always work), ura 4+, his3, leu1 or sup3-5 markers should be used. The latter marker is an opal nonsense suppressing tRNA gene which suppresses ade6-704. This marker has a deleterious effect for the cell when present in several copies. On minimal medium supplemented with 10 µg/ml adenine or yeast extract medium ade6-704 mutant colonies are red but when suppressed by sup3-5 they are white. If a sup3-5 containing plasmid is not integrated into the genome then instability leads to cells lacking sup3-5 and hence to the formation of pink colonies. This contrasts with clones containing one copy of the integrated plasmid which are white, and enables a rapid distinction to be made between integrated and non integrated clones. Alternatively, ade6 itself can be transformed into an ade 6 deleted strain. For ura 4+ plasmids the best strain to use is ura 4-D18 that contains a complete deletion of the S. pombe ura 4+ gene and integration by homologous recombination at the ura 4 locus is thus avoided. Recently, the kanMX6 module has been widely used as a heterologous dominant marker that allows selection of G418-resistant cells in S.pombe (see Gene Disruption and Replacement Method II).

1. Transform a yeast strain with the plasmid of interest.

2. Isolate a transformant colony and grow up in 100 ml of YES medium (i.e. non selective conditions) for about 20 generations (re-inoculate 1ml of this culture into 100 ml of fresh YES medium 2-3 times).

3. Plate out about 1000 cells/plate onto selective medium and incubate until colonies form. These colonies should be stable due to integration of the plasmid into the genome. This can be tested by replica plating to YES medium twice and then back to selective medium.

4. Confirm the integration by Southern blotting

1. Transform with the plasmid and plate out in the absence of adenine.

2. Replica plate to minmal plates with 10µg / ml adenine.

3. After 2-3 days look for white fast growing colonies which are putative integrants. The pink colonies still have free plasmid.

1. Make a disruption of the gene of interest by inserting the ura 4+ marker or the LEU2 marker gene in the ORF. If possible delete as much of the ORF as possible and conserve at least 1kb either side.

2. Purify this linear fragment.

It is then possible to delete the native gene by integrating the marker flanked by flanking sequences of the gene of interest. This integration is performed by transformation of a diploid strain, as below. Homologous integration with a double crossover will result in excision of the native gene.

3. Transform either a h-/h+ura 4-D18/ura 4-D18 ade6-M210/ade6-M216 or a h-/h+leu1-32/leu1-32/ ade6-M210/ade6-M216 diploid strain, depending on the marker used, with 100ng, 300ng, 1µg of the fragment (less DNA is necessary if transforming using the Okayama or electroporation methods) . Select diploid transformants expressing the ura 4+ or LEU2 gene.

Gene replacements in S. pombe can be carried out using the same approach. A gene replacement event can be selected using a diploid strain in which one copy of the gene is disrupted with the ura 4+ gene, because ura 4- cells generated when the disrupted gene is replaced will be resistant to the drug 5-fluororotic acid (5-FOA). Thus, replacement of the ura 4+ disrupted gene with a linear DNA fragment containing the in vitro altered gene will convert the cells to a 5-FOA-resistant phenotype. It is necessary to maintain the presence of 5-FOA in the medium, as the effect is reversible. 5-FOA is used at 1 mg / ml in plates. NB - this is an expensive chemical. It is always wise to check for the occurrence of the event by Southern blotting of the genomic DNA of the new construct.

1. Design primers: 100mer oligos with 80nt to gene of interest and 20nt to ura4/kan module; primers should be HPLC purified.

2. Do 2-5 independent PCR reactions.

Pool reactions

Purify the DNA by phenol/chloroform extraction and concentrate by ethanol precipitation. Resuspend in 10µl of TE.

3. Follow lithium acetate method II (see section 2.1) for transforming PCR products, with the following modifications:

Grow cells in YE5S instead of MM.

Plate on YE5S with NO kanamycin.

Incubate overnight at 30°C.

Replica plate the resultant lawn onto YE5S with 100µg/ml kanamycin.

Incubate at 30°C.

You will have large and small colonies- pick the large ones and restreak, etc.

1. Take transformant colony and streak out to single colonies on YES agar with no selection for about 3 days until colonies form.

2. Replica plate to selective medium e.g. 35°C on YES for a ts strain , 25°C on minimal for the auxotrophic marker and check for the stability of the prototrophic (and / or other plasmid-borne) phenotypes. The sup3-5/ade6-704 system (described above) is particularly useful for this purpose.

1. Construct a plasmid containing a selectable marker (LEU 2) and the wild type copy of the entire chromosomal region of interest.

2. Digest the plasmid with a restriction enzyme to completely remove the ORF. Purify the linear fragment containing the plasmid with the upstream and downstream flanking DNA sequences .

3. Transform 1 µg of this fragment into the strain containing the allele of interest. Identify transformants expressing the selectable marker. The gap in the plasmid is repaired using the mutant chromosomal sequences as a template.

4. Recover the plasmid from yeast as described below.

The efficiency of recovery of "repaired" plasmids versus "recircularised" plasmids appears to depend on the size of the flanking sequences, 1kb either side is recommended. On average 25 % of the plasmids obtained are "repaired". If overexpression of the mutant allele is deleterious to the cell it may not be possible to recover the repaired plasmid.

Plasmid recovery from S. pombe is difficult as 2 µm based plasmids often seem to form multimers, and rearrangements (tandem duplication or deletions) are frequent. This problem can be avoided by using the plasmid pFL20 or derivatives that contain the stb element and remain as monomers. Furthermore, ars1 plasmids, which comprise many of those currently in use, do not tend to rearrange and are easy to recover. It is worth noting also that yeast material in the final preparation appears to inhibit E. coli transformation and thus using more of the preparation may not be a good idea To recover a plasmid from a transformant there are two procedures which are routinely used:

2. Spin down the cells 3000 rpm 5 minutes.

3. Resuspend in 1.5 ml:

50mM Citrate/ Phosphate pH 5.6 ( 7.1 g/l Na2HPO4, 11.5 g/l citric acid)

.2M Sorbitol

(Adjust to pH 5.6 with 5M NaOH.)

2 mg/ml Zymolyase-20T (added after autoclaving).

transfer to an Eppendorf tube and incubate at 37°C for 1hour.

4. Pellet the cells in an Eppendorf centrifuge for 30 seconds. Resuspend in 300 µl TE.

5. Add 35 µl 10% SDS, mix and incubate at 65°C for 5 minutes.

6. Add 100 µl 5M potassium acetate, mix and leave on ice for 30 minutes.

7. Spin down at 4°C for 10 minutes.

8. Add 50 µl of supernatant to 100 µl NaI solution (Geneclean Kit, Stratech Scientific Ltd.) with 5 µl glassmilk (Geneclean Kit, Stratech Scientific Ltd.).

9. Incubate for 5 minutes at room temperature.

10. Spin for 5 seconds (maximum) at room temperature, discard the supernatant and wash the pellet three times with 400 µl of ice-cold NEW wash (Geneclean Kit, Stratech Scientific Ltd.).

11. Elute DNA twice with 10 µl of TE at 55°C for 3 minutes each time.

12. Spin out the glass milk and keep the supernatant.

13. Transform 5 µl of the supernatant into 100 µl of competent E.coli JA226 cells.

The use of the Geneclean or Qiagen kits improves the transformation frequency by at least 10 fold; also it is very important to use a recBC E.coli strain such as JA226 when using 2 µm or non-ars containing plasmids. For ars1 based plasmids recA strains like E. coli DH5 can be used.

A second method is also used; this has been less well tried in this lab but is quicker and reportedly gives good results, as follows:

B. 1. Grow small cultures (at least 1.4 ml) with selection.

2. Collect the cells by a 5 second spin in a microfuge.

3. Decant away the supernatant and briefly vortex the tube to resuspend the pellet in the residual medium.

4. Add 0.2 ml of

2 % Triton X-100

1 % SDS

100mM NaCl

10mM Tris HCl (pH 8.0)

5. Add 0.2 ml phenol:chloroform:IAA and 0.3 g acid washed glass beads.

6. Vortex for 2 minutes and then microfuge for 5 minutes.

7. Take upper aqueous layer to a fresh Eppendorf and extract with 200µl phenol:chloroform:IAA.

8. Precipitate the DNA , wash with 70% EtOH, dry and resuspend in 10µl TE.

9. Use 1-5 µl for transformations of competent E.coli or yeast .

Note: This method also works well for isolating genomic DNA.

The standard Thiamine concentration for repression of gene expression from the nmt-1,-41 and -81 promoters is 15µM (200µl per 400ml medium of a 10mg/ml stock solution stored in the dark at 4°C), although 2µM Thiamine is usually sufficient for repression of the promoter in most cases, and may aid induction. Certain "toxic" gene products require extra suppression and growth can be aided by increasing the concentration up to 60µM especially when using nmt1 which is the strongest and leakiest of these promoters. Despite the above, it is not really possible to partially induce the promoter in a population by using low thiamine concentrations.

A Thiamine concentration of 2nM in minimal medium plates is sufficient for cells overexpressing prep3x-nmt1-rum1, to grow to a colony size of 0.1-1mm in three days at 32°C, before running out of Thiamine. Such colonies consist of cells showing the rum1 overexpression phenotype (which ran out of Thiamine before running out of nutrition - at the border of the colony) and of starved wild-type cells (which ran out of nutrition before running out of Thiamine - in the centre of the colony). Although the rum1 overexpression phenotype is lethal the strains can be recovered very easily by transferring the starved cells to +Thiamine medium. This method has been used in a screen designed to isolate cDNAs whose products alter cell morphology when overexpressed. It has the advantage that no replica plating from +Thiamine to -Thiamine plates is required.

The minimal Thiamine concentration mentioned above might have to be increased or further decreased to give similar results when overexpressing other genes and/or when using another promoter.

Promoter strength (and leakiness): nmt1 > nmt41 > nmt81

If a strain is used for a screen containing ectopic expression of a gene from the nmt promoter, mutations frequently affect the activity of the nmt promoter. These should be screened out in a secondary screen, e.g. by Northern Analysis.

S. pombe cells coordinate cell growth and the cell cycle. Exponentially growing wildtype cells are born at a similar cell size and double their mass before entering the next round of cell division. This is the result of a size control coupling growth and the cell cycle. They grow by tip elongation, therefore cell length is a measure of the cell cycle stage of a cell. For this reason if you size select cells for instance by elutriation, they will all be at the same stage in the cell cycle.

To achieve this, exponentially growing wildtype S. pombe cells use a size control in G2 which controls the size requirement for entry into mitosis. However, if this size control is eliminated by mutation or an environmental change, then a cryptic G1 size control comes into play. Rapidly growing wildtype cells are born at a cell size that exceeds this G1 size control so it does not delay S-phase. However, small S. pombe cells spend a larger portion of the cell cycle in G1 and use this control to coordinate growth and cell division. The existence of a size control, means that blocking cell growth leads to cell cycle arrest.

Cell number and O.D. will increase exponentially and in step with each other. In smaller cells such as wee mutants, the same will be true but cells will now spend a greater proportion of their time in G1 as visible by FACS analysis.

In fact both small cells and large cells have the same generation time: the time for one cell to double its mass to become two cells. This means that the O.D. and cell number will increase at the same rate. However for the same O.D., populations of large cells will contain less cells than populations of small cells. So a small cell accumulates less actual mass than a large cell in the same time, but the small cell's percentage increase in mass is the same as that of the large cell.

The coupling of cell mass increase and cell division can be uncoupled by certain mutations and environmental conditions. Under conditions of nitrogen starvation for instance, wildtype cells undergo mitosis at a reduced cell size and complete several rounds of the cell cycle, with little increase in cell mass, and as a result become smaller. This leads to an increase in the proportion of cells in G1, as the G1 size control comes into play. This effect can be accentuated by the addition of pheromone to responsive cells.

Cells becoming smaller.

Cell number increasing to a greater extent than the O.D..

A larger G1 peak will be seen by FACS analysis.

Cell cycle blocks also uncouple growth from cell division, this is how they were defined. If the cell division cycle is blocked, then cells continue to accumulate mass.

Cells will become longer.

O.D. will increase exponentially (until cells lyze).

Cell number will remain constant. (see transition point for discussion of the time at which cell number will stop increasing).

Re-replicating cells will show increasing DNA content.

There are two accurate means of determining cell number, a haemocytometer or electronic cell counters.

Samples are fixed in formal saline and then the number of cells in a fixed volume is counted (together with the mean cell volume) as they pass into a probe with either a 70µm or a 100µm aperture. The fluid circulating through the Coulter counter is ISOTON.

1. Fix cells by adding 400µl of culture to 1.6mls formal saline (0.9% saline, 3.7% formaldehyde) in a 30ml Universal tube. Vortex and store at 4°C (samples keep for months).

2. Dilute with an appropriate volume of ISOTON. You want to aim to have roughly 1-3 x 105 cells in each 500µl aliquot that is counted, to ensure that the count is well above background. The machine cannot count greater than 6 x 105 cells / 500µl aliquot. The exact number is not crucial and typically you can dilute your 2mls of fixed cells by adding 18mls ISOTON (assuming the samples were from a mid-log culture).

3. Preparing the Coulter counter:

- Turn on (3 switches including the mean cell volume unit)

- Set 'Corrected Count' to Auto

- Set 'Current' to 200

- Set 'Full Scale' to 10 mA

- Set 'Polarity' to Auto

- Set 'Lower Threshold' to 070 (for cells) or 050 (for spores)

- Set 'Upper Threshold' to 999

- Set 'Alarm Threshold' to OFF

- Set 'Attenuation' to 8 (100µm probe) or 16 (70µm probe)

- Set 'Preset Gain' to 4 (100µm probe) or 2 (70µm probe)

4. Open the door and insert a Universal containing ISOTON. Focus the aperture - you will be able to see fluid moving in. If there is anything blocking the hole then simply wipe it with you finger. You now need to clean out the probe (and so bring the background down) by taking a series of 'counts', each of which will wash ISOTON through the system. To do this turn the 'Reset/Count' knob clockwise to 'Count'. At the end of each count you will get a reading of the background counts. Turn the knob to 'Reset' and continue the washes (only ever turning the knob in a clockwise direction) until the background is down to about 300 or less. If you have problems then try the 'Fill / Drain' knob once and continue the 'counts'.

5. Once the background is down then you're ready to go. Each sample should be sonicated (30-60 seconds on setting 6) before counting. Count each sample 3 times and take the mean. Note the mean cell volume. It is important to take each count at the same position (the 'Reset / Count' knob has 2 count positions - either can be used but stick to the same one for all the counts). It is important to keep an eye on the probe during the counts (you'll be able to see it) to ensure that the aperture is clear. If it blocks then simply wipe it clean and repeat the count.

6. Once you've finished, put a Universal containing ISOTON back in the machine and repeat the washing counts until the background is low once again. Immerse the probe in the ISOTON bath and switch off.

Notes :

- If you have a series of samples that relate to each other (e.g. a time course) it is important to count them all at a single sitting. Counts taken on different occasions (when the background will be different) are not directly comparable.

- Don't let the probe dry out at any time - always keep it immersed in ISOTON.

- Calculating cell no. / ml : for 2mls fixed cells containing 0.4mls culture, diluted with 18mls ISOTON, multilply mean count by 100 to get the actual cells / ml of your culture (remember that the mean count represents 500µl of your 20mls of diluted cells).

1) Check the waste is empty if not empty it.

2) Switch the machine on and wait for the front page to appear

3) Put your sample into the plastic container and put onto the WBC probe, press Count

4) The number of cells/ml will be displayed on the screen. To get the actual cell number multiply by 100 (assuming that you have 400µl of sample in 20ml), if you want a printout press Print. Do two

counts for each sample.

5) When you have finished clean the probe 3x with Isoton by pressing Count. Leave the probe in Isoton and press Clean for 10sec. Switch off still pressing Clean.

6) Empty the waste.

1) Very long cells will not be counted. If your cells are very elongated then you should use the Coulter Counter.

2) If the machine becomes clogged it will automatically recount. If it remains clogged then try cleaning it (press Clean for a few secs) and Flush and Fill. If it is still clogged then repeat everything, if you still cannot unclog it then go and have a cup of tea and try again later.

3) Nitrogen starved cells or spores may not count accurately as the lower discriminator which is set automatically, is in the wrong

place (I don’t know why). To change it go to the Menu page and press 4 (Manual Discrimination) and enter, press 1(WBC) and enter. You should now be on the Manual Discrimination page. Press enter to select the lower discriminator which will become a solid line

when it is selected, move it with the arrow keys and then press

enter and the new reading will be displayed. The upper discriminator does NOT affect the cell number, it is used to discriminate between different populations. To go back to the Front page press Select (X3). Unfortunately you have to reset the lower discriminator for each new sample.

1. Scan picture of the cell(s) containing a calibration bar taken at the same resolution.

2. Open the programme NIH Image 1.59 ppc and import image(s).

3. Use broken line in Tools Box and make a line from one end of the cell to the antipodal end.

4. Use "measurement" command under "Analyse" in Menu to measure cell length; measurements are stored and can be visualised by "Show Results" command; measurements can be undone with "Undo Measurement" under "Edit" Menu..

4. Calibrate by measuring calibration bar in a likewise fashion.

The execution point of a cell cycle gene is that point in the cell cycle at which the gene function is required. One way of estimating the execution point of a gene is to measure the transition point of a conditional mutant of that gene. The transition point of a particular mutant is that point in the cell cycle at which a defective gene product will block progression through the cell division cycle.

In an asynchronously growing culture shifted to restrictive conditions, only those cells at points in the cycle beyond the transition point will go on to divide before blocking at the transition point in the next cycle.

The transition point is therefore calculated from the fractional increase in cell number when an asynchronous culture is shifted to the restrictive condition. It is given by the following formula :

T.P. = 1 - ln (1+A) T.P. = transition point

ln 2 A = fractional increase in cell number

The transition point can more simply be estimated as follows :

In the S. pombe life cycle septation occurs at about the same time as the peak in S phase. This means that for a G1 / S mutant even those cells before the transition point will go on to divide once when an asynchronous culture is shifted to restrictive conditions. Some cells in the culture will divide twice, those after the transition point but after the peak of septation, so the ratio of final cell number to the initial value will be greater than 2. This means that transition points of G1 / S mutants can be negative. Cells complete G1 in the cell division cycle preceding the one in which they will complete the subsequent G2.

1. In order to be confident that the transition point actually reflects the execution point of your gene it is necessary to show that the same value is obtained with a number of alleles. Differing values can result from such factors as a particular allele taking a long time to inactivate at the restrictive temperature, or being partially inactive even at the permissive temperature.

2. If your gene has more than one execution point in the cell cycle the transition point will only reflect the latest one with respect to septation.

Caution: If too many cells are loaded on to the rotor they starve during elutriation and do not recover synchronously. If there are dead cells in the starting culture these will come off with the small cells!

Cells are eluted into conditioned media into which some fresh media has been added, which should be kept at the elutriation temperature after it is generated by loading cells onto the rotor. The first few hundred mls of culture should then be discarded as this usually contains many dead cells.

After elutriation cells take up to 1 hour to recover, before this the cells are not growing exponentially. The first peak of septation occurs approximately within the next hour, depending on the temperature at which point sampling should begin. The septation index is usually monitored every 20 minutes and should reach a maximum of about 20% and a minimum of 0%. Cells can be followed for up to 2 cell cycles. As the culture will be growing and dividing, progressively smaller samples will need to be taken if each sample is to contain the same number of cells.

Alternatively cell cycle mutants can be used to block the cell cycle at a particular point. On return to the permissive temperature cells will then proceed synchronously through the subsequent phases of the cycle. Leaky mutants are not suitable for this method, some mutants do not recover well on return to the permissive temperature. This approach has been used successfully to study mitotic onset (via cdc25-22 block and release experiments) and can also be used to study exit from mitosis (Yanagida's lab uses dis2 block and release - nda3 can also be used). cdc10-129 can be used to study passage through 'Start' and S phase, however cells are slow to enter the subsequent mitosis. Such experiments have the disadvantage of being rather unphysiological, due to the effects of blocking the cell cycle for at least one generation.

11mM hydroxyurea is used to arrest cells at the beginning of S-phase. As most cells in an exponentially growing culture are in G2 it takes about a generation time to arrest the population with HU. The arrest is not maintained for long and cells will typically begin to leak through the block after about 4 - 5 hours, depending on the temperature and medium (leak through is faster in rich medium and at higher temperatures). It is possible to add a second dose of 11mM HU after 4 hours which will hold cells in the block for several more hours. Release from hydroxyurea by 2 washes in HU free media enables cells to synchronously pass through S-phase. The subsequent mitosis however lacks synchrony, and occurs only 1.5 hours after the release. This suggests a DNA damage checkpoint may come into play.

Unlike S. cerevisiae it is not possible to block S. pombe in mitosis with microtubule binding drugs.

Thiabendazole is very effective at depolymerising microtubules in S. pombe but does not cause a good c