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CHAPTER 13
Yeast
INTRODUCTION
Saccharomyces cerevisiae (baker’s yeast) and Schizosaccharomyces pombe (fission yeast)
are often considered to be model eukaryotic organisms, in a manner analogous to
Escherichia coli as a model prokaryotic organism. Both yeasts have been extensively
characterized and their genomes completely sequenced. They are as easy to grow as
other microorganisms, and they have a haploid nuclear DNA content only 3.5 times that
of E. coli. However, despite the small genomes sizes, these yeasts display most of the
features of higher eukaryotes. The fact that many cellular processes are conserved among
different eukaryotic species-combined with the powerful genetic and molecular tools that
are available- has made these yeasts important experimental organisms for a variety of
basic problems in eukaryotic molecular biology.
Primarily for historical reasons, most studies on yeast have involved Saccharomyces
cerevisiae (hereafter termed yeast). Culturing yeast is simple, economical, and rapid,
characterized by a doubling time of ∼90 min on rich medium. In addition, yeast has been
well adapted to both aerobic and anaerobic large-scale culture. Cells divide mitotically
by forming a bud, which pinches off to form a daughter cell. The progression through the
cell cycle can be monitored by the size of the bud; this has been used to isolate a large
collection of mutants (called cdc mutants) that are blocked at various stages of the cell
cycle. Since yeast can be grown on a completely defined medium (see UNIT 13.1), many
nutritional auxotrophs have been isolated. This has not only permitted the analysis of
complex metabolic pathways but has also provided a large number of mutations useful
for genetic analysis. Mutations can be generated by classical UV or EMS mutagenesis
(UNIT 13.3B) or by transposon mutatgenesis (UNIT 13.3), which allows one to identify the
mutation via the inserted transposon.
Yeast can exist stably in either haploid or diploid states. A haploid cell can be either of
two mating types, called a and α. Diploid a/α cells—formed by fusion of an α cell and
an a cell (UNIT 13.2)—can grow mitotically indefinitely, but under conditions of carbon
and nitrogen starvation will undergo meiosis. The meiotic products, called spores, are
contained in a structure called an ascus. After gentle enzymatic digestion of the thick cell
wall of the ascus, the haploid spore products can be individually isolated and analyzed
(UNIT 13.2). This ability to recover all four products of meiosis has allowed detailed
genetic studies of recombination and gene conversion that are not possible in most other
eukaryotic organisms. The existence of stable haploid and diploid states also facilitates
classical mutational analysis, such as complementation tests and identification of both
dominant and recessive mutations.
The haploid yeast cell has a genome size of about 15 megabases and contains 16 linear
chromosomes, ranging in size from 200 to 2200 kb. Thus, the largest yeast chromosome
is still 100 times smaller than the average mammalian chromosome. This small chromosome size, combined with the advent of techniques for cloning yeast genes as well
as manipulating yeast chromosomes, has allowed detailed studies of chromosome structure. Three types of structural elements required for yeast chromosome function have
been identified and cloned: origins of replication (ARS elements), centromeres (CEN
elements), and telomeres. The cloning of these elements has led to the construction of
Yeast
Current Protocols in Molecular Biology 13.0.1-13.0.4, April 2008
Published online April 2008 in Wiley Interscience (www.interscience.wiley.com).
DOI: 10.1002/0471142727.mb1300s82
C 2008 John Wiley & Sons, Inc.
Copyright 13.0.1
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artificial chromosomes that can be used to study various aspects of chromosome behavior, such as how chromosomes pair and segregate from each other during mitosis and
meiosis. In addition, systems using artificial chromosomes have been designed that allow
cloning of larger contiguous segments of DNA (up to 400 kb) than are obtainable in other
cloning systems. These structural elements, as well as cloned selectable yeast genes, have
permitted the construction of yeast/E. coli shuttle vectors that can be maintained in yeast
as well as in E. coli (UNITS 13.4 & 13.6).
Procedures for high-efficiency transformation of yeast (UNIT 13.7) have been available for
nearly two decades, allowing cloning of genes by genetic complementation (UNITS 13.8
& 13.9). Because yeast has a highly efficient recombination system, DNAs with alterations in cloned genes can be reintroduced into the chromosome at the corresponding
homologous sites (UNIT 13.10). This has permitted the rapid identification of the phenotypic consequences of a mutation in any cloned gene, a technique generally unavailable
in higher eukaryotes. In addition, homologous recombination permits a wide variety of
genetic techniques that have greatly facilitated the analysis of biological processes.
Despite its small genome size, yeast is a characteristic eukaryote, containing all the
major membrane-bound subcellular organelles found in higher eukaryotes, as well as
a cytoskeleton. Yeast DNA is found within a nucleus and nucleosome organization of
chromosomal DNA is similar to that of higher eukaryotes, although no histone H1 is
present. Three different RNA polymerases transcribe yeast DNA, and yeast mRNAs
(transcribed by polymerase II) show characteristic modifications of eukaryotic mRNAs
[such as a 5 methyl-G cap and a 3 poly(A) tail], although only a few S. cerevisiae
genes contain introns. Transcriptional regulation has been extensively studied and at
least one yeast transcriptional activator has been shown to function in higher eukaryotes
as well. High-molecular-weight yeast DNA and RNA can be prepared fairly quickly
(UNITS 13.11 & 13.12). Another characteristic of eukaryotes is the proteolytic processing
of precursor proteins to yield functional products, which is often coupled to secretion.
Yeast has several well-studied examples of secreted proteins and pheromones, and the
large number of genes that have been identified as involved in protease processing and
secretion suggests a highly complex pathway. Yeast protein extracts can be prepared
using three different protocols (UNIT 13.13); the best choice will depend on the particular
application. The ease and power of genetic manipulation in yeast facilitate the use of this
organism to detect novel interacting proteins using the two-hybrid system or interaction
trap (UNIT 20.1).
Although Saccharomyces cerevisiae is the most commonly studied yeast, S. pombe is
also an important experimental organism (UNIT 13.14). Although both yeasts are unicellular
microorganisms that grow in similar medium, they are evolutionarily quite distant. It has
become increasingly clear that, in terms of molecular mechanisms, S. pombe is more
similar to higher eukaryotic organisms than S. cerevisiae. Experimental manipulations
in S. pombe are broadly similar to those in S. cerevisiae, although the technical details
often differ. The chapter includes units on S. pombe relating to strain maintenance and
media (UNIT 13.15), growth and genetic manipulation (UNIT 13.16), and introduction of DNA
into cells (UNIT 13.17).
This chapter is written for the molecular biologist who has not previously worked with
yeast. The glossary below introduces the terms of yeast molecular biology.
aerobic growth growth in the presence of oxygen, utilizing the Krebs cycle.
Introduction
α and a factor mating type–specific polypeptides secreted by either α or a haploid cells,
respectively, which interact with haploid cells of the opposite mating type to stimulate
mating.
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anaerobic growth growth in the absence of oxygen, utilizing fermentation (via glycolysis).
ARS elements DNA sequences present throughout the yeast genome that confer autonomous replication on plasmids in yeast; most of these sequences also function as
chromosomal origins of replication as well.
ascus thick-walled sac containing the four haploid products, called spores, resulting
from meiosis.
cdc mutants strains of yeast that exhibit stage-specific blocks of the cell cycle; these
mutations define genes important in DNA replication, meiosis, and sporulation.
CEN element DNA sequences present at the centromere that ensure proper segregation
of chromosomes during mitosis and meiosis, presumably by promoting interaction with
the mitotic spindle.
cir+ strains of yeast that contain the naturally occurring 2µm plasmid.
ciro yeast strains that have lost this endogenous 2µm plasmid.
δ element ∼330-bp sequence, present as direct repeats at the ends of the transposable
element Ty1, and also found dispersed throughout the genome.
gene disruption a mutation constructed in vitro in a cloned gene which, upon reintroduction into the genome at the homologous chromosomal site, results in inactivation of
the gene function.
glusulase a digestive enzyme isolated from snails that breaks down thick cell walls of
either an ascus to allow isolation of spore products, or a yeast cell to produce spheroplasts.
heterothallic common laboratory strains of yeast which—due to a mutation in the HO
gene—stably maintain a given allele at the MAT locus.
homothallic strains of yeast (typically found in the wild) that, in a haploid state, rapidly
interconvert the MAT locus, resulting in rapid switching between the α and a mating
types; cultures of such strains rapidly diploidize.
killer strains strains of yeast that harbor a double-stranded RNA virus; such strains kill
sensitive yeast strains via secretion of a protein toxin, to which killer strains are immune.
MAT the mating-type locus which is expressed and therefore determines the mating type
of a haploid cell; this locus has two alleles—the MATa allele confers the a mating type,
while MATα specifies the α mating type.
meiosis the process by which the number of chromosomes present in a diploid cell is
halved to yield haploid products.
mitosis vegetative cell division (of either haploid or diploid cells) in which the chromosome number stays the same.
petites mutants of yeast (either nuclear or mitochondrial) with impaired mitochondria
function; they grow as small colonies on fermentable carbon sources and are unable to
grow on nonfermentable carbon sources.
schmoo a distinctive shape of a haploid cell (pear-shaped), induced by exposure to
mating pheromone.
Yeast
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sporulation the end product of meiosis, induced by carbon and nitrogen starvation of a
diploid cell, which results in four haploid progeny contained as spores within an ascus;
this complicated developmental process requires over 200 genes.
telomeres DNA sequences found at the end of linear chromosomes that are essential
for chromosome stability and complete replication; in S. cerevisiae, telomeres consist of
tandem repeats of the sequence 5 dG1-3 dT3 .
Ty1 elements the primary transposable element found in yeast, which functions as a
retrotransposon (via reverse transcription of its RNA and subsequent reinsertion into the
genome).
UAS element upstream activating sequences in yeast promoters, to which regulatory
proteins bind in order to enhance the rate of transcription.
zygote a morphologically distinct cellular structure formed by the fusion of two haploid
cells of opposite mating type, which results in formation of a diploid.
Zymolyase β-glucanase, isolated from Arthrobacter luteus, that hydrolyzes the yeast
cell wall and is used to prepare spheroplasts for a variety of purposes.
KEY REFERENCES
Watson, J.D., Hopkins, N.H., Roberts, J.W., Steitz, J.A., and Weiner, A.M. 1987. Yeasts as the E. coli of
eukaryotic cells. In Molecular Biology of the Gene, Vol. 1, pp. 550-594. Benjamin/Cummings, Menlo
Park, Calif.
Strathern, J.N., Jones, E.W., and Broach, J.R. (eds.) 1981. The Molecular Biology of the Yeast Saccharomyces: Metabolism and Gene Expression Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
Strathern, J.N., Jones, E.W. and Broach, J.R. (eds.) 1982. The Molecular Biology of the Yeast Saccharomyces:
Metabolism and Gene Expression. Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
Victoria Lundblad and Kevin Struhl
Introduction
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BASIC TECHNIQUES OF YEAST GENETICS
SECTION I
Like Escherichia coli, yeast can be grown in either liquid media or on the surface of (or
embedded in) solid agar plates. Yeast cells grow well on a minimal medium containing
dextrose (glucose) as a carbon source and salts which supply nitrogen, phosphorus, and
trace metals. Yeast cells grow much more rapidly, however, in the presence of protein and
yeast cell extract hydrolysates, which provide amino acids, nucleotide precursors, vitamins, and other metabolites which the cells would normally synthesize de novo. During
exponential or log-phase growth, yeast cells divide every 90 min when grown in such
media. Log phase can be divided into three stages based on the rate of cell division (or
the proportion of budded cells within a culture), which is in turn a function of the cell
density of the culture. As cell density increases, nutrient supplies drop and the rate of cell
division slows (the measurement of cell density, as well as techniques for the propagation
and genetic manipulation of yeast, are described in UNIT 13.2). Early-log phase is the period
when cell densities are <107 cells/ml. Mid-log phase cultures have densities between 1
and 5 × 107 cells/ml. Late-log phase occurs when cell densities are between 5 × 107 and
2 × 108 cells/ml. At a density of 2 × 108 cells/ml yeast cultures are said to be saturated
and the cells enter stationary, or G0 phase.
Recipes for media that are commonly encountered when working with yeast are provided
in UNIT 13.1. The rich medium, YPD (yeast extract, peptone, dextrose; also called YEPD
media), is most commonly used for growing S. cerevisiae. Additional recipes are provided
for minimal and complete minimal dropout media, which are routinely used for testing
mating type, selecting diploids, determining auxotrophic requirements (UNIT 13.2), and
selecting for transformants (UNIT 13.7). Finally, recipes for media required in the more
advanced techniques described in UNITS 13.6 and 13.10 are presented.
Preparation of Yeast Media
UNIT 13.1
Preparation of sterile media of consistently high quality is essential for the genetic
manipulation of yeast. Recipes for media needed in the protocols in this chapter are
provided below. It is recommended that ingredients always be purchased from the same
manufacturer. The following sources are recommended for specific ingredients (complete
addresses and phone numbers are provided in APPENDIX 4):
J.T. Baker
dextrose
ammonium sulfate
potassium acetate
glycerol
Difco
agar (Bacto-agar)
peptone (Bacto-peptone)
yeast extract (Bacto-yeast extract)
yeast nitrogen base (YNB, without
amino acids or ammonium
sulfate)
Sigma
amino acids
nucleotide bases
canavanine
cycloheximide
L-α-aminoadipic acid
galactose (with 0.01%
contaminating glucose)
potato starch
PCR, Inc
5-fluoroorotic acid
Autoclaving is usually carried out for 15 min at 15 lb/in2, but times should be increased
when large amounts of media are being prepared (20 min for 4 to 6 liters and 25 min for
6 to 12 liters).
Contributed by Douglas A. Treco and Victoria Lundblad
Current Protocols in Molecular Biology (1993) 13.1.1-13.1.7
Copyright © 2000 by John Wiley & Sons, Inc.
Saccharomyces
cerevisiae
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Supplement 23
LIQUID MEDIA
Ingredients for liquid media are dissolved in water to 1 liter, mixed until completely
dissolved, and autoclaved in 100- or 500-ml media bottles. Alternatively, liquid media
can be filter sterilized, resulting in faster preparation, less carmelization (of dextrose),
and faster growth of cells. Recipes for “premixes” are provided to minimize the number
of materials that must be weighed each time media is prepared. When preparing premixes,
break up any large chunks of dextrose before mixing with other components, then shake
the container vigourously until contents are homogenized. It is convenient to make the
premix in the empty plastic containers in which 2.5 kg of dextrose is packaged. Throughout this chapter, YNB −AA/AS refers to yeast nitrogen base without amino acids or
ammonium sulfate. (See listing of recommended suppliers of ingredients in unit introduction above.)
Rich Medium
YPD medium
Per liter:
10 g yeast extract
20 g peptone
20 g dextrose
Premix:
250 g yeast extract
500 g peptone
500 g dextrose
Use 50 g/liter
Final concentration:
1% yeast extract
2% peptone
2% dextrose
This rich, complex medium—also known as YEPD medium—is widely used for the growth
of yeast when special conditions are not required.
It is preferable to use a 20% (10×) solution of dextrose that has been filter sterilized or
autoclaved separately (and added to the other ingredients after autoclaving) to prevent
darkening of the media and to promote optimal growth.
Minimal Media
Minimal medium
Per liter:
1.7 g YNB −AA/AS
5 g (NH4)2SO4
20 g dextrose
Premix:
68 g YNB −AA/AS
200 g (NH4)2SO4
800 g dextrose
Use 27 g/liter
Final concentration:
0.17% YNB −AA/AS
0.5% (NH4)2SO4
2% dextrose
This minimal medium—also known as synthetic dextrose (SD) medium—can support the
growth of yeast which have no nutritional requirements. However, it is used most often as a
basal medium to which other supplements are added (see CM dropout medium below).
Complete minimal (CM) dropout medium, per liter:
1.3 g dropout powder (Table 13.1.1)
1.7 g YNB −AA/AS
5 g (NH4)2SO4
20 g dextrose
(Alternatively, replace last three ingredients with 27 g minimal medium premix)
Preparation of
Yeast Media
CM dropout powder, also known as minus or omission powder, lacks a single nutrient but
contains the other nutrients listed in Table 13.1.1. Complete minimal (CM) dropout medium
is used to test for genes involved in biosynthetic pathways and to select for gene function in
transformation experiments. To test for a gene involved in histidine biosynthesis one would
determine if the yeast strain in question can grow on CM minus histidine (−His) or “histidine
dropout” plates. It is convenient to make several dropout powders, each lacking a single
nutrient, to avoid weighing each component separately for all the different dropout plates
required in the laboratory.
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Table 13.1.1
Nutrient Concentrations for Dropout Powdersa
Nutrientb
Adenine (hemisulfate salt)
L-arginine (HCl)
L-aspartic acide
L-glutamic acid (monosodium salt)
L-histidine
L-leucinef
L-lysine (mono-HCl)
L-methionine
L-phenylalanine
L-serine
L-threoninee
L-tryptophan
L-tyrosine
L-valine
Uracil
Amount in
dropout
powder(g)c
2.5
1.2
6.0
6.0
1.2
3.6
1.8
1.2
3.0
22.5
12.0
2.4
1.8
9.0
1.2
Final conc.
Liquid
in prepared
stock conc.
media (µg/ml) (mg/100 ml)d
40
20
100
100
20
60
30
20
50
375
200
40
30
150
20
500
240
1200
1200
240
720
360
240
600
4500
2400
480
180f
1800
240
aCM dropout powder lacks a single nutrient but contains the other nutrients listed in this table. Nomenclature
in this manual refers to, e.g., a preparation that omits histidine as histidine dropout powder. Conditions from
Sherman et al., 1979.
bAmino acids not listed here can be added to a final concentration of 40 µg/ml (40 mg/liter).
cGrind powders into a homogeneous mixture with a clean, dry mortar and pestle. Store in a clean, dry bottle
or a covered flask.
dUse 8.3 ml/liter of each stock for special nutritional requirements. Store adenine, aspartic acid, glutamic acid,
leucine, phenylalanine, tyrosine, and uracil solutions at room temperature. All others should be stored at 4°C.
eWhile these amino acids can be used reliably when included in autoclaved media, they supplement growth
better when added after autoclaving.
fUse 16.6 ml/liter for L-tyrosine nutritional requirement.
It may be preferable to use a 10× solution of dropout powder (i.e., 13 g of dropout powder
in 100 ml water) that has been “sterilized” separately (and added to the other ingredients
after autoclaving) to improve the growth rate in this medium.
Sporulation medium, per liter
10 g potassium acetate (1% final)
1 g yeast extract (0.1% final)
0.5 g dextrose (0.05% final)
This nitrogen-deficient “starvation” medium contains acetate as a carbon source to promote
high levels of respiration, which induces diploid yeast strains to sporulate. Sporulation can
be carried out in liquid media or on plates (see below and UNIT 13.2). If nutrients are required,
add them at the concentrations listed in Table 13.1.1.
Alternative Carbon Sources
Wild-type yeast can use a variety of carbon sources other than glucose to support growth.
These include galactose, maltose, fructose, and raffinose. In particular, galactose is often
used to induce transcription of sequences fused to the GAL10 promoter (UNIT 13.6). All are
used at a concentration of 2% w/v (20 g/liter) and should be made by replacing dextrose
in the recipe for minimal or complete minimal (CM) dropout media. For a nonfermentable
carbon source—which will not support the growth of petites (cells lacking functional
mitochondria; see glossary)—2% potassium acetate (w/v), 3% glycerol, 3% ethanol, or
2% glycerol and 2% ethanol (v/v each) can be used. YPA medium (2% potassium acetate,
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2% peptone, and 1% yeast extract) is excellent for inducing high levels of respiration in
cells prior to sporulation (UNIT 13.2).
SOLID MEDIA
Making solid media for yeast is—for the most part—no different from preparing plates
for bacteriological work (see UNIT 1.1). For all plates, agar is added at a concentration of
2% (20 g/liter). A pellet of sodium hydroxide (∼0.1 g) should be added per liter to raise
the pH enough to prevent agar breakdown during autoclaving. In addition, add a stir bar
to facilitate mixing after autoclaving.
After autoclaving, flasks are left for 45 to 60 min at room temperature until cooled to 50°
to 60°C. (Drugs and other nutrients are added after 30 min at room temperature.) Just
prior to pouring, put the flask on a stir plate at medium to high speed and mix until contents
are homogeneous (∼5 min). After pouring, a few bubbles can be removed from the agar
surface by passing the flame of a Bunsen burner lightly over the surface of the molten
agar (“flaming” the plates). One liter of media will yield 30 to 35 plates.
While a specific brand of petri plate is not required, we recommend Fisher plates (100 ×
15 mm, #8-757-12), which have ridges around the tops of the covers to allow easy
stacking, making plate pouring less cumbersome.
When preparing the plate recipes below, follow the general guidelines for mixing and
autoclaving in the introduction to liquid media (p. 13.1.2). Most plates can be stored at
room temperature for ≤4 months. Plates containing drugs (cycloheximide, 5-fluoroorotic,
canavanine, and L-α-aminoadipic acid) or Xgal are stable for 2 to 3 months when stored
at 4°C.
Minimal Plates and Rich Plates
YPD, minimal, and CM dropout plates
Per liter: Follow recipes for liquid media above, adding 20 g agar and a pellet of
NaOH.
Premixes: Follow recipes for liquid media premixes above, adding 500 g agar for
YPD premix and 800 g agar for minimal premix. To prepare plates, add one NaOH
pellet and the following amounts of premix (per liter):
YPD plates—70 g YPD plate premix
Minimal plates—47 g minimal plate premix
CM dropout plates—47 g minimal plate premix + 1.3 g dropout powder
Specialty Plates
The recipes for α-aminoadipate, canavanine, and cycloheximide plates are included even
though no specific use for them is described in this chapter. They are commonly used in
negative selection experiments in the same way that 5-FOA plates are used (see below).
One can select against the wild-type LYS2, CAN1, and CYH2 genes by growth on plates
that select for cycloheximide, α-aminoadipate, or canavanine resistance, respectively (for
review, see Brown and Szostak, 1983).
Preparation of
Yeast Media
5-fluoroorotic acid (5-FOA) plates
To a 2-liter flask (containing a stir bar), add:
1 g 5-FOA powder
500 ml H2O
5 ml 2.4 mg/ml uracil solution
Stir with low heat ∼1 hr until completely dissolved; filter sterilize
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To a separate 2-liter flask, add:
1.7 g YNB −AA/AS
5 g (NH4)2SO4
20 g dextrose
20 g agar
1.3 g uracil dropout premix (Table 13.1.1)
H2O to 500 ml and autoclave
(Alternatively, replace first four ingredients with 47 g minimal plate premix)
When the molten agar cools to ∼65°C, gently add the sterile 5-FOA/uracil solution
to the uracil dropout medium by pouring it down the inside wall of the flask
containing the latter. Swirl gently to mix and pour the plates.
URA3+ strains are unable to grow on media containing the pyrimidine analog 5-fluoroorotic
acid (Boeke et al., 1984). This observation has led to methods that use 5-FOA to select against
the functional URA3 gene (see UNIT 13.10). This type of selection—termed “negative selection” (Brown and Szostak, 1983)—can also be used to select against the wild-type LYS2,
CAN1, and CYH2 genes.
5-FOA is quite expensive, and plates should be used sparingly. The material is prepared in
bulk and is substantially discounted for members of the Genetics Society of America
(Bethesda, Md.).
Xgal plates, per liter
1.7 g YNB −AA/AS
5 g (NH4)2SO4
20 g dextrose
20 g agar
0.8 g dropout powder (omitting appropriate amino acids; see Table 13.1.1)
NaOH pellet
(Alternatively, replace first four ingredients with 47 g minimal plate premix)
Add H2O to 900 ml and autoclave. Add 100 ml of 0.7 M potassium phosphate, pH
7.0, and 2 ml of 20 mg/ml Xgal prepared in 100% N,N-dimethylformamide (stored
as frozen stock; see Table 1.4.2).
Dissection plates
Follow the recipe for YPD plates, keeping in mind that dissection plates used for
tetrad analysis should be of uniform thickness and free of imperfections. After the
plates have been poured they should be flamed. Stack the plates on a level surface
in piles of six and move gently in a circular motion to “even out” the agar. Certain
batches of agar produce plates that have microscopic precipitates embedded in the
agar, often looking much like yeast spores. If this occurs, an agar of higher purity
can be used, such as Noble agar (Difco) or agarose.
α-aminoadipate plates
1.7 g YNB −AA/AS
20 g dextrose
20 g agar
H2O to 1 liter
Add ingredients to a 2-liter flask and autoclave. When the molten agar cools to
∼65°C, add 34 ml of a solution of 6% L-α-aminoadipic acid (prepared by dissolving
α-aminoadipate in 100 ml water and adjusting the pH to ∼6.0 with 1 M KOH). The
final concentration of α-aminoadipic acid in the plates should be 0.2%. Swirl gently
to mix and pour the plates.
Lys2− yeast use α-aminoadipic acid as an alternate nitrogen source. Since yeast can use
certain amino acids as nitrogen sources, only those amino acids which are required by the
particular strain being used should be added to these plates. These can be added from liquid
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stocks prior to autoclaving. When using this media for isolating lys2− yeast, lysine must be
added at 30 ìg/ml.
Canavanine plates
Follow recipe for complete minimal (CM) dropout plates, omitting the nutrient
arginine. When the 1-liter autoclaved solution has cooled to ∼65°C, add 10 ml of 6
mg/ml filter-sterilized canavanine sulfate solution. The final concentration of canavanine should be 60 µg/ml.
The sterile canavanine sulfate solution can be stored frozen.
Cycloheximide plates
Follow the recipe for YPD plates. When the agar cools to ∼65°C, add 1 ml of 10
mg/ml filter-sterilized cycloheximide solution. The final concentration of cycloheximide in the plates should be 10 µg/ml.
The sterile cycloheximide stock solution can be stored frozen.
STRAIN STORAGE AND REVIVAL
Yeast strains can be stored at −70°C in 15% glycerol (viable for >5 years), or at 4°C on
slants consisting of rich medium supplemented with potato starch (viable for 1 to 2 years).
Both methods are described below.
BASIC
PROTOCOL
Preparation and Inoculation of Frozen Stocks
Make a solution of 30% (w/v) glycerol. Pipet 1 ml into 15 × 45-mm, 4-ml screwcap vials.
Loosely cap the vials and autoclave 15 min.
To inoculate vials for storage, add 1 ml of a late-log or early-stationary phase culture,
mix, and set on dry ice. Store at −70°C. Revive by scraping some of the cells off the frozen
surface and streak onto plates. Do not thaw the entire vial. Cells can also be stored in the
same way by adding 80 µl dimethyl sulfoxide (DMSO) to 1 ml cells (8% v/v) and storing
at −70°C.
ALTERNATE
PROTOCOL
Preparation and Inoculation of Slants
1. For 250 slants, add the following ingredients to a 1-liter flask:
70 g YPD plate premix
20 mg adenine (hemisulfate salt)
20 g potato starch
H2O to 500 ml
Excess adenine prevents ade− mutations from being lost.
2. Set the flask in a 4-liter beaker filled with 1 liter of water. Place the beaker on a
heat-controlled, magnetic stirring apparatus and stir with the heat setting on high.
With smooth and continuous stirring, the contents should not burn and should be molten
after ∼1 hr.
3. With the entire setup in place, begin pipetting 2-ml aliquots into 15 × 45-mm, 4-ml
screwcap vials. When all vials have been filled, put the caps on loosely, pack in the
original boxes, and autoclave 15 min.
4. Lean the boxes against a support at an angle of ∼70°. Allow the slants to dry 2 days
and screw the caps on tightly.
Preparation of
Yeast Media
Slants can be stored at room temperature for at least 6 months.
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5. To inoculate a slant, smear cells from the flat end of a sterile toothpick onto the agar
surface of the slant. Cap loosely and incubate 1 or 2 days at 30°C. After growth, screw
the cap on as tightly as possible and store at 4°C.
Slants are a convenient way to store and mail strains. They can be mailed immediately after
inoculating since sufficient growth will occur in transit. See another method for mailing
strains below.
Mailing and Reviving Strains
Yeast strains can be conveniently mailed as slants. Alternatively, transfer cells to a piece
of sterile Whatman 3MM paper by pressing the paper onto the desired yeast colony using
forceps that have been dipped in ethanol and flamed. Wrap the paper in sterile aluminum
foil and mail to recipient.
BASIC
PROTOCOL
Revive the strain by placing the paper (yeast side down) on the surface of an agar plate.
Incubate the plate at 30°C. A thick patch of yeast should be visible after lifting the paper.
LITERATURE CITED
Boeke, J.D., LaCroute, F., and Fink, G.R. 1984. A positive selection for mutants lacking orotidine-5′-phosphate decarboxylase activity in yeast: 5-fluoroorotic acid resistance. Mol. Gen. Genet. 197:345-346.
Brown, P.A. and Szostak, J.W. 1983. Yeast vectors with negative selection. Meth. Enzymol. 101:278-290.
Sherman, F., Fink, G.R., and Lawrence, C.W. 1979. Methods in Yeast Genetics. Cold Spring Harbor
Laboratory, Cold Spring Harbor, N.Y.
Contributed by Douglas A. Treco
Massachusetts General Hospital
Boston, Massachusetts
Victoria Lundblad
Baylor College of Medicine
Houston, Texas
Saccharomyces
cerevisiae
13.1.7
Current Protocols in Molecular Biology
Supplement 19
Growth and Manipulation of Yeast
1
UNIT 13.2
2
Douglas A. Treco and Fred Winston
1
2
Massachusetts General Hospital, Boston, Massachusetts
Harvard Medical School, Boston, Massachusetts
ABSTRACT
Yeast cultures can be grown, maintained, and stored in liquid media or on agar plates using
techniques similar to those for bacterial cultures. This unit describes culture conditions
for these basic techniques. Additional methods describe determination of yeast mating
type, diploid construction, sporulation, tetrad dissection, and random spore analysis.
C 2008 by John Wiley & Sons, Inc.
Curr. Protoc. Mol. Biol. 82:13.2.1-13.2.12. Keywords: yeast r media r yeast culture r growth
INTRODUCTION
This unit first presents the necessary details for growing yeast cells (see Basic Protocols
1 to 3). This is followed by a description of replica-plating methods for assessing the
nutritional requirements (see Basic Protocol 4) and mating type (see Basic Protocol
5) of strains. Yeast genetic experiments often require the construction of strains with
specific genotypes, as well as an analysis of the meiotic segregation patterns of newly
introduced mutations (see UNIT 13.8). These genetic manipulations are carried out using
the protocols presented in the final sections of this unit, which describe the construction
and selection of diploids, sporulation, and tetrad analysis (see Basic Protocols 6 to 8).
The protocols in this unit are for use with the budding yeast Saccharomyces cerevisiae.
Different growth media and protocols are used for another yeast, Schizosaccharomyces
pombe (UNITS 13.14-13.17).
Aside from different media requirements, yeast cells are physically manipulated essentially as described for bacterial cells—i.e., they are grown in liquid culture (in tubes or
flasks) or on the surface of agar plates, and are manipulated using the basic equipment
described in UNITS 1.1-1.3. In addition, a well-equipped yeast laboratory requires static
and shaking incubators dedicated to 30◦ C and a microscope with magnification up to
400×. A second microscope adapted for dissecting yeast tetrads is extremely valuable
for the genetic analyses and strain constructions described in this unit. A small electric
clothes dryer is indispensable when replica plating is done frequently and large numbers
of velvets are regularly used.
Because meiosis and sporulation are parts of the life cycle of S. cerevisiae, it is relatively
straightforward to create strains with different genotypes. Genes on different chromosomes sort independently, and linked genes can be separated by recombination. Diploids
are constructed from parents that will each contribute some of the markers desired in
the haploid products. The approach for constructing a new yeast strain is presented in
several distinct stages: (1) diploid construction, where two haploid strains are mated
(see Basic Protocol 6); (2) sporulation, where diploid cells are induced to form spores
(see Basic Protocol 7); (3) tetrad preparation, where the ascus wall is removed from the
tetrad (see Basic Protocol 8); and (4) tetrad dissection, where each of the four haploid
spores from a single tetrad is specifically positioned on a plate and grown for subsequent
studies (see Basic Protocol 8). Not all of these steps are necessary in an individual experiment. Although in many cases yeast strains can be constructed by transformation using
gene replacement methods (UNIT 13.10), strain construction by crosses and tetrad analysis
is still a widely used and important method in yeast genetics.
Current Protocols in Molecular Biology 13.2.1-13.2.12, April 2008
Published online April 2008 in Wiley Interscience (www.interscience.wiley.com).
DOI: 10.1002/0471142727.mb1302s82
C 2008 John Wiley & Sons, Inc.
Copyright Yeast
13.2.1
Supplement 82
BASIC
PROTOCOL 1
GROWTH IN LIQUID MEDIA
Wild-type S. cerevisiae grows well at 30◦ C with good aeration and with glucose as a
carbon source. When using culture tubes, vortex the contents briefly after inoculation to
disperse the cells. Erlenmeyer flasks work well for growing larger liquid cultures, and
baffled-bottom flasks to increase aeration are especially good. It is important that all
glassware be detergent-free.
For good aeration, the medium should constitute no more than one-fifth of the total
flask volume, and growth should be carried out in a shaking incubator at 300 rpm. For
small-scale preparations of DNA and RNA, yeast can be grown in glass or plastic culture
tubes filled one-third full with medium and shaken at 350 rpm in a rack firmly attached
to a shaking incubator platform.
BASIC
PROTOCOL 2
BASIC
PROTOCOL 3
GROWTH ON SOLID MEDIA
Yeast cells can be streaked or spread on plates as shown for bacteria in the sketches in
UNIT 1.3. When a dilute suspension of wild-type haploid yeast cells is spread over the
surface of a YPD plate and incubated at 30◦ C, single colonies may be seen after ∼24 hr
but require ≥48 hr before they can be picked or replica plated (see below). Growth on
dropout media (UNIT 13.1) is about 50% slower.
DETERMINATION OF CELL DENSITY
The density of cells in a yeast culture is most reliably determined by direct counting in a
hemacytometer chamber (UNIT 1.2) and plating for viable colonies (UNIT 1.3). The density of
a culture can also be determined spectrophotometrically by measuring its optical density
at 600 nm (OD600 ). However, before this can be a reliable method, it is necessary to
calibrate the spectrophotometer by graphing the OD600 as a function of cell density as
determined by direct counting and plating for viable colonies (titering). Different mutant
backgrounds can affect the cell size or shape, thereby altering the OD/cell. For reliable
measurements by OD, cultures should be diluted so that the OD600 is <1. For wild-type
yeast strains in this range, 0.1 OD600 units correspond to ∼3 × 106 cells/ml. Thus, an
OD600 of 1 is equal to ∼3 × 107 cells/ml.
Some mutant strains have a “clumpy” phenotype, where cells are stuck together. These
clumps will result in inaccurate density measurements. Therefore, in such strains it
is important to disperse the clumps by mild sonication prior to counting, plating, or
measuring the OD.
BASIC
PROTOCOL 4
Growth and
Manipulation of
Yeast
DETERMINATION OF PHENOTYPE BY REPLICA PLATING
Cells from yeast colonies grown on any medium can be tested for their nutritional
requirements by replica plating (UNIT 1.3). An inexpensive replica plating block can be
constructed by gluing a circular plexiglass disk (8-cm diameter, 1 cm thick) onto the end
of a hollow Plexiglas tube (8 cm long with an 8-cm outer diameter). Sterile velveteen
squares (velvets) are held in place by a large adjustable tube clamp (available in any
automotive supply outlet) set to fit snugly around the outside of the tube.
A master plate containing the strain or strains of interest is first printed onto a velvet.
A copy of this impression is transferred to plates made with all the relevant selective
media, which may include various dropout and drug media, as well as alternative carbon
sources (UNIT 13.1). For analysis of temperature-sensitive mutations (UNIT 13.8), a copy of
the master plate is made on a plate that will provide all the nutritional requirements of
the strain. This plate is then incubated at 37◦ C. For any replica plating tests, a permissive
plate should be the last plate used, as a positive control for transfer of the yeast cells.
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DETERMINATION OF MATING TYPE
Genetic analysis of yeast requires knowledge of the mating type. This protocol is based
on the ability of a strain with a single auxotrophic requirement (the tester strain) to
complement any and all nutritional requirements of strains of the opposite mating
type, as long as the genetic deficiency in the tester strain is not present in any of the
uncharacterized strains. The genetic deficiency found in the tester strain prevents it from
growing on minimal plates. This deficiency is complemented by the wild-type gene in
the uncharacterized strains, which themselves usually cannot grow on minimal plates
due to one or more auxotrophic mutations. When strains of opposite mating type mate,
the resulting diploid can grow on minimal plates. The following protocol is useful for
determining the mating types of the numerous spore colonies produced during strain
construction.
BASIC
PROTOCOL 5
Materials
S. cerevisiae: MATα thr4− (tester), MATα thr4− (tester), and uncharacterized
strains
YPD medium and plates (UNIT 13.1)
Minimal plates (UNIT 13.1)
Replica plating block (UNIT 1.3)
Sterile velvets (UNIT 1.3)
1. Grow 1-ml overnight cultures of each tester strain.
2. Spread 200 µl of each tester strain on a YPD plate. To get a completely even
distribution of cells, replica plate this freshly spread YPD plate onto a sterile velvet,
lift and rotate 90◦ , and print again. Lift, rotate, and print once more. Discard velvet
and incubate plate overnight at 30◦ C.
3. On the next day, replica plate the strain to be tested onto two fresh YPD plates.
Discard velvet. Onto one of these two YPD plates, replica plate one of the two tester
strains prepared in step 2. Using a new velvet, repeat with the other YPD plate and
the other tester strain.
If many uncharacterized strains are picked into a grid pattern on a single plate (UNIT 1.3),
their mating types can be determined simultaneously.
4. Incubate plates ≥4 hr at 30◦ C.
5. Replica plate each plate onto a minimal medium plate. Incubate plates overnight at
30◦ C.
6. Score for mating type.
Growth on the minimal plate printed with the MATa thr4− tester strain indicates that the
uncharacterized strain is MATα. Growth on the minimal plate printed with the MATα
thr4− tester strain indicates that the uncharacterized strain is MATa.
The thr4− mutation makes tester strains auxotrophic for threonine (i.e., they can’t grow
on threonine dropout medium). However, a mutation in any gene that leads to nutritional
auxotrophy can be used for the tester strains, and strains with defects in other genes
required for threonine biosynthesis can be tested using MATa or MATα thr4− testers.
One or a few strains can be tested simply by patch-mating the uncharacterized cells to each
tester strain (see Basic Protocol 6) for 4 hr and streaking each pair onto minimal plates.
Strains without auxotrophic requirements cannot be tested using the above protocol for
mating-type determination. If such a strain arose from one of four spores from a tetrad,
the mating type can be inferred if it can be determined for the other three strains from
the same tetrad. Alternatively, the mating type can be determined by observing zygote
formation microscopically. Patch-mate the strains of unknown mating type separately to
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MATa and MATα strains. After 4 hr at 30◦ C, examine the cells microscopically for zygote
formation (zygotes will appear as relatively large 2- and 3-lobed structures in which the
lobes are connected by wide, smooth “necks”). If the strain with unknown mating type was
haploid, zygote formation will only be obvious in one of the two mixtures. The unknown
mating type is opposite that of the strain with which it forms zygotes.
Mating type can be determined in ∼24 hr if fresh tester plates are available. A stock of
tester plates can be prepared as described in step 1 and stored at 4◦ C for 1 to 2 months.
Allow mating (step 4) to proceed overnight when using tester plates that have been stored
for more than a few days.
BASIC
PROTOCOL 6
DIPLOID CONSTRUCTION
Diploids are constructed by mating strains of opposite mating types on the surface of
agar plates (patch mating). Mix cells from freshly grown colonies of each haploid parent
with a toothpick in a circle ∼0.5 cm in diameter on an agar plate (the plate should allow
growth of both haploid strains). Allow mating to proceed ≥4 hr at 30◦ C, then streak the
mating mixture onto a plate that will select for the diploid genotype. Diploids can be
constructed and stored indefinitely.
When there is no selection specific for the diploid genotype (the case when one of the
haploid parents has the same nutritional requirements as the diploid), isolate diploids by
physically “pulling zygotes” out of the mating mixture using a dissecting microscope.
After mating for 4 hr, transfer a small dab of cells to a YPD plate, making several
parallel lines with the toothpick. The cells will be diluted in each streak. Using the
dissecting microscope, identify zygotes by their characteristic shape (described in Basic
Protocol 5, step 6), pick them up with the dissecting needle, move them away from
the streak of cells, and set them down. To ensure that the selected cells are actually
diploids, patch them onto sporulation plates (UNIT 13.1) and examine microscopically for
tetrad formation after appropriate incubation (see following protocols). Alternatively,
attempt to mate selected cells with a pair of mating type tester strains and examine microscopically for zygote formation with each tester. Zygotes should not form
if a diploid was correctly selected.
BASIC
PROTOCOL 7
SPORULATION OF DIPLOID CELLS
Starvation of diploid yeast cells for nitrogen and carbon sources induces meiosis and spore
formation, during which chromosomes replicate and proceed through two divisions to
produce haploid nuclei. These nuclei (along with surrounding cytoplasm) are individually
packaged into spores, and the four spore products of a single meiosis (tetrad) are held
together in a thick-walled sac (ascus).
The sporulation process can be induced in cells growing on solid or in liquid medium.
For unknown reasons, some strains do not sporulate well on plates. Even for strains that
do, the efficiency can often be increased by sporulation in liquid. Because some strains do
not sporulate well on plates and others do not sporulate well in liquid, both methods are
presented here. One of the two methods should result in reasonably good spore formation
for any given diploid. Spores can be stored at 4◦ C for 1 to 2 weeks without a significant
decrease in viability.
Materials
Growth and
Manipulation of
Yeast
Yeast cells
Sporulation plates or sporulation medium, with appropriate nutrients (UNIT 13.1)
YPD medium (UNIT 13.1)
13.2.4
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Sporulation on plates
1a. Patch cells that have been grown on YPD or selective plates onto a sporulation plate.
If no selective conditions are required, grow cells several days on YPD plates prior to
transfer to sporulation medium. Allow single colonies to grow for 3 to 4 days on YPD;
for patches of cells, allow 2 days growth on YPD. While this pregrowth is not essential, it
results in much more efficient sporulation. A small dab of cells should be smeared over a
relatively large area (∼1 cm2 ) of the sporulation plate, such that no thick patches of the
inoculum are visible.
2a. Incubate 4 days at 25◦ C.
Sporulation is generally less efficient at higher temperatures. While sporulation can occur
in the absence of amino acids or other nutrients that are required by the strain for mitotic
growth, sporulation is much more complete when those (and only those) nutrients that
the particular strain requires are added (see Table 13.1.1).
Several rounds of mitotic growth will cause the cell number to visibly increase over the
sporulation incubation period.
3a. Visualize tetrads by suspending a small dab of cells in a drop of water on a microscope
slide and examining at a magnification of 250× to 400×.
Tetrads will appear as clusters of four small spheres (the spores), all held within a tightfitting sac (ascus). The four spores can be in either a diamond or tetrahedral configuration.
Not all asci will contain four spores. Some cells do not package all four spore products.
The proportion of cells that undergo sporulation as well as the fraction of four-spored
asci that result varies from strain to strain.
Sporulation in liquid media
1b. Pick a single colony of the diploid and grow it overnight in YPD.
2b. Inoculate 3 ml of fresh YPD with the overnight culture, diluting the cells ∼50-fold.
3b. Grow the culture at 30◦ C until the cells are at a concentration of 1–2 × 107 cells/ml.
4b. Centrifuge the cells 5 min at 1200 × g and wash twice with sterile water.
5b. Resuspend the cells in 2 ml liquid sporulation medium and transfer to a small glass
tube.
Liquid sporulation medium contains 1% (w/v) potassium acetate.
6b. Add any required amino acids to the sporulation medium and incubate the tube with
shaking for 3 days at room temperature.
For many strains, the cultures can be moved to 30◦ C after the first day to speed up
sporulation.
If a strain appears refractory to induction of sporulation, try pregrowing cells in YPA
medium (UNIT 13.1). The use of acetate as a carbon source requires respiration, which is
a requirement for sporulation. This requirement for respiratory competence is reflected
by the fact that cells that have lost mitochondrial function (petites; see glossary in
introduction to this chapter) are unable to sporulate.
PREPARATION AND DISSECTION OF TETRADS
The analysis of yeast tetrads requires a standard light microscope with a stage that is
movable along both the x and y axes in precisely measurable intervals, but that does
not move up and down (focusing is accomplished by moving the objectives). The
microscope must be modified with an assembly for mounting an inverted petri dish
on its stage and a micromanipulator for holding and moving a fine glass needle (see
Support Protocol for preparation of dissecting needles). Plans for the construction of
BASIC
PROTOCOL 8
Yeast
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Supplement 82
micromanipulators for tetrad analysis have been published (Sherman, 2002). An assembled instrument is available from at least three vendors: MVI (http://www.mvi-inc.
com/TDM400.htm), Zeiss (http://www.zeiss.com/4125681f004ca025/Contents-Frame/
6b6194aae761e001c1256cbf0039b80c), and Singer Instruments (http://www.singerinst.
co.uk/test/).
Needles used for tetrad dissection can be made in the laboratory from 2-mm glass rods
(see below). However, it is much easier, although more expensive, to purchase dissecting
needles from Singer Instruments. As an alternative, needles can be made from 0.002-in.
(∼50-µm) fiber-optic glass (Sherman, 2002).
To prepare yeast tetrads for dissection, the asci are first incubated in a dilute solution of
Zymolyase-100T to break down the ascus cell wall. An older method, using glusulase,
although relatively inexpensive, is no longer widely used, as Zymolyase-100T generally
gives more uniform and reproducible digestion of the ascus wall.
Dissection of yeast tetrads requires a steady hand and a great deal of patience. Don’t be
discouraged, as anyone can learn to be good at it. It is not uncommon for a skilled yeast
geneticist to dissect up to 60 tetrads per hr. However, dissection of 20 to 30 tetrads per
hr is a commendable and highly feasible goal.
Materials
Spores (see Basic Protocol 7)
0.5 mg/ml Zymolyase-100T (ICN Immunobiologicals) in 1 M sorbitol
Dissecting microscope
Dissecting needle (see Support Protocol)
Prepare the tetrads
1a. For spores from a plate: Add a small toothpick-full of tetrads to 50 µl of 0.5 mg/ml
Zymolyase-100T solution and gently resuspend.
1b. For spores from liquid cultures: Microcentrifuge 1 ml cells for 10 sec. Pour off
supernatant, and resuspend pellet in 50 µl of 0.5 mg/ml Zymolyase-100T solution.
2. Examine cells under a light microscope to detect intact asci.
3. Incubate 10 min at 30◦ C.
4. Very gently add 0.8 ml sterile water by slowly running it down side of the tube.
After Zymolyase treatment, the four spores should remain associated, but more loosely
than when initially examined. The wall of the ascus should be expanded and loosely
associated with the spores.
5. Set tubes on ice and leave them there. Streak treated spores (using an inoculating
loop, a glass pipet, or an automatic pipettor with tip) in two parallel lines across the
surface of a YPD dissection plate (UNIT 13.1), as shown in Figure 13.2.1.
6. Examine the plate, inverted with the lid off, on the dissecting microscope.
Individual tetrads, grouped into tetrahedral or diamond-shaped clusters of spores, should
be visible.
Dissect the tetrads
The following procedure is specifically adapted for the dissecting microscope described
by Sherman (2002; Fig. 13.2.1).
Growth and
Manipulation of
Yeast
7. Position the plate so that streaks are parallel to the x axis of the stage. Focus on
spores and position a good tetrad in the center of the field.
13.2.6
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Figure 13.2.1 Essential features of the dissection microscope. Not all parts are shown, including
the light source and condenser (below) and the objective and eyepiece (above). The micromanipulator is attached to the base of the microscope. The mark with the large dot on the y-axis guide is
aligned with demarcations on the right side of the stage to identify positions a, b, c, and d (corresponding to positions 10, 15, 20, and 25 respectively, in this figure). The upper right-hand corner
of the plate holder is aligned with demarcation along the rear of the stage to identify positions 1,
2. . .13 (corresponding to positions 60, 65. . .120, respectively in this figure).
At 100× magnification, attempt to dissect a tetrad that is not near any other tetrads, so
that multiple spores from unrelated tetrads are not picked up by the needle.
Be careful not to graze the tip of the dissecting needle with the petri dish as it is being placed on or removed from the stage. Lower the needle and rotate the microscope
objectives away from the stage before changing plates.
8. Move dissecting needle upward using coarse adjustment to a position such that its
tip is touching the plate surface when the joystick is depressed halfway.
Moving the joystick down results in the needle moving upward. When the joystick is fully
up, the needle will be visible as a large black sphere looming below the plate surface. It
will appear to be much larger than the tetrad itself.
9. Using a downward motion on the joystick, gently touch needle to surface of plate,
immediately adjacent to the tetrad.
The tetrad should disappear under the needle. When attempting to pick up individual
spores or tetrads, touch the needle to the plate very close to, but not directly on top of,
the spore or tetrad.
10. Gently lift the joystick straight up.
The tetrad should appear to be gone. At this point the tetrad should be on the tip of
the needle. If it is still on the plate, repeat steps 9 and 10 until the spores stick to the
needle. Pick up tetrads or individual spores as gently as possible, with the needle barely
touching the agar surface. If Zymolyase treatment was effective, all four spores should
move together. Tetrads will fall apart at this stage if the treatment was excessive.
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11. Using stage movement controls, move the plate so that the needle is at rightmost
position on the plate (using x-axis control) and at a position (position a in Fig. 13.2.1)
∼1 cm away from streak of treated spores (using y-axis control).
The four spores will be set in a line perpendicular to the original streak at positions a,
b, c and d. These positions should be 0.5 cm apart and are easily found by aligning an
arbitrary marking on the movable stage with a fixed y-axis scale that is even with the
right side of the stage (Fig. 13.2.1).
12. Use the joystick to gently touch the needle to the agar surface.Then use the joystick
to move the needle down and forward, so that the point where the needle touched
the agar is visible.
The tetrad should be on the agar surface. The following variations are possible:
a. No spores were deposited. In this case repeat step 12, using more force when touching
the needle to the agar.
b. Only a single spore is on the plate (the best result). Move the y-axis adjustment to
position b and repeat step 12 to deposit the next spore.
c. If two or three spores are on the plate, move the y-axis adjustment to position b and
attempt to set down the remaining one or two spores on the needle. If two spores
were on the needle and only one spore is deposited at position b, move the needle to
position c and set down the remaining spore. If two spores were deposited at position
b, the needle must be used to break them apart. This is best accomplished by moving
the joystick in circular motion, so that the tip of the needle hits the plate and drags
over the spores. When the pair of spores is broken apart, pick one up and move it to
position c. Don’t forget that there are multiple spores at position a that still need to
be separated.
d. If all four spores are present, use the needle to break them apart. Once this is done,
pick up one or more of the spores (leaving one at position a) and attempt to lay down
a single spore at each of the three remaining positions.
Be careful not to move the x-axis controls during the dissection of an individual tetrad.
If the four spores cannot be separated after repeatedly dragging with the needle, the
zymolyase treatment was probably not sufficient. Let the treated culture (which should
have been on ice) set at room temperature for a few minutes and then streak onto a new
YPD dissection plate.
13. Use x- and y-axis controls to move the plate back to a position where the needle is
directly below the streak of treated spores. Repeat steps 7 through 12, with spores
of each successive tetrad set down in a line 0.5 cm to right of and parallel to that of
the preceding tetrad.
Placement of these lines is determined by aligning an arbitrary marking on the movable
stage (upper right-hand corner of the plate in Fig. 13.2.1) with a fixed x-axis scale that is
even with back edge of stage. Alternatively, a strip of paper marked in 0.5-cm intervals
can be taped along front edge of stage. In this case the lower right-hand corner of the
plate holder is used as the arbitrary alignment point.
14. Continue steps 7 through 13 until all positions along the x axis are occupied by
dissected tetrads.
The plate is then rotated 180◦ for dissection of additional tetrads using the streak on the opposite side of plate as a source. Thirteen tetrads can be dissected on each side of the plate.
SUPPORT
PROTOCOL
Growth and
Manipulation of
Yeast
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PREPARATION OF DISSECTING NEEDLES
One of the most difficult steps involved in tetrad analysis is the preparation of the
dissecting needle. Like tetrad dissection, making needles requires patience and practice.
The general strategy is to first produce a long fine thread of glass, which is broken into
multiple short segments. Individual segments are then glued to the ends of capillary
tubes. The finished product is L-shaped, with a short arm of ∼1.3 cm, culminating in a
needle “tip” that is 40 to 150 µm in diameter (see Fig. 13.2.2).
Current Protocols in Molecular Biology
Figure 13.2.2
Preparation of glass needles for tetrad dissection.
1. Create a thin glass thread by heating a piece of glass tubing until it is flexible, and
then, in one motion, remove it from the heat source and pull the ends apart very
quickly.
The goal here is to draw out the molten glass into a very fine thread before it (nearly
instantly) hardens. Timing is everything here, so a few attempts may be necessary before
a thread that is suitably thin can be generated. Either capillary tubes or Pyrex glass
pipets can be used as sources of glass tubing. Suitable threads should be between 40 and
150 µm in diameter, depending upon application and personal taste. Human hairs, which
range from ∼40 to 100 µm in diameter, can be used as a reference.
2. Once a thread of appropriate diameter is obtained, break it into segments ∼1.3 cm
long. To do this, simply begin at one end, pulling the segments straight away from
the rest of the thread using your fingers. Wear latex gloves for a better grip.
Alternatively, segments can be cut using a fresh razor blade.
3. Examine the segments microscopically. Look for chips and protrusions in the glass
as shown in Figure 13.2.2. Save any segments with an acceptable end, noting which
end is which, and discard those with chipped or uneven ends.
The best needles (those that easily pick up and release cells and do not cut the surface of
the agar) have an absolutely flat surface that is perpendicular to the shaft (see Fig. 13.2.2).
4. Create a dissecting needle by attaching a segment with an acceptable tip to the end
of a capillary tube. Introduce a right-angle bend in the capillary tube 0.5 to 1.0 cm
from the end by briefly heating the tube in a Bunsen burner flame and bending the
tip using forceps. Glue the glass thread in place, either inside the short arm of the
L-shaped capillary tube or against the outside edge, using any fast-acting glue (see
Fig. 13.2.2).
The finished needle is L-shaped with a short arm ∼1.3 cm long culminating in the flat
working surface.
5. Examine the finished product after mounting the needle on a dissecting microscope.
Place an agar plate on the microscope and press the needle firmly against the surface
of the agar to make sure the needle leaves an even, circular impression.
Even with all these precautions, needles will perform differently when it is time to dissect,
so several candidates should be made at the same time and tested to determine which
function best.
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ALTERNATE
PROTOCOL
RANDOM SPORE ANALYSIS
As an alternative to separating spores by tetrad dissection, meiotic products can be
released from their asci, dispersed by sonication, and plated directly onto agar plates.
The spore colonies can then be screened for the desired genotypes by replica plating.
This method is generally used only when large numbers of spores are needed. Tetrad
analysis provides much more information than random spore analysis, as the four products
of a single meiosis are analyzed.
Materials
Spores (see Basic Protocol 7)
1 mg/ml Zymolyase-20T (ICN Immunobiologicals) in H2 O, filter sterilized
2-Mercaptoethanol (2-ME)
1.5% (v/v) Nonidet P-40 (NP-40)
Ethanol
Sonicator and probe
Additional reagents and equipment for cell counting (UNIT 1.2) and replica plating
(UNIT 1.3)
Prepare the tetrads
1a. For spores from a plate: Resuspend several toothpicks-full of tetrads in 50 ml water
in a 50 ml flask.
1b. For spores from liquid cultures: Microcentrifuge 1 ml sporulation culture for 10 sec.
Pour off supernatant, and resuspend pellet in 5 ml water.
2. Add 0.5 ml of 1 mg/ml Zymolyase-20T solution and 10 µl of 2-ME.
Lyse unsporulated cells
3. Incubate overnight at 30◦ C with gentle shaking.
Treatment of the sporulated culture with Zymolyase-20T in a hypotonic solution results in
lysis of unsporulated diploid cells. The preparation should be examined microscopically
after the Zymolyase treatment to evaluate its effectiveness. A higher concentration of the
enzyme or the more concentrated preparation of Zymolyase (Zymolyase-100T) can be
used.
4. Add 5 ml of 1.5% Nonidet P-40 (NP-40). Transfer the suspension to a 15-ml disposable tube and set 15 min on ice.
5. Hold the tube in one hand and insert the sonicator probe as far into the liquid as
possible, but without touching the bottom or the sides of the tube. Before sonicating,
clean the sonicator probe with water followed by a wipe-down with ethanol.
Sonicate, plate, and analyze spores
6. Sonicate 30 sec at 50% to 75% full power, then set on ice 2 min. Repeat twice.
Sonication produces heat that will warm up the spore suspension significantly. The tube
should be cooled between the sonication steps.
7. Centrifuge spores 10 min at 1200 × g. Aspirate or pour off supernatant and resuspend
in 5 ml of 1.5% NP-40. Vortex vigorously. Repeat twice.
8. Sonicate as in step 6 (with repeats).
Growth and
Manipulation of
Yeast
The spores should be examined after the last sonication step to ensure that no spores
remain stuck together. More sonication steps at higher power settings will release the
more tenacious spores. If spores remain stuck to each other, add 2 ml glass beads (Type I,
Sigma) and shake 30 min at 300 rpm in an Erlenmyer flask at 30◦ C. Let the beads settle
and remove the supernatant containing the spores.
13.2.10
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Current Protocols in Molecular Biology
9. Centrifuge spores 10 min at 1200 × g. Aspirate or pour off supernatant and resuspend
in 5 ml water. Vortex vigorously. Repeat.
10. Count a 10-fold dilution of the treated spores using a hemacytometer.
11. Dilute the spores with water to get 103 spores/ml. Plate 100 µl on several YPD plates
and incubate 3 days at 30◦ C.
12. Screen spore colonies for markers of interest by replica plating (see Basic Protocol
4 and UNIT 1.3).
COMMENTARY
Background Information
Diploids are constructed from parents that
each contribute some desired markers in the
haploid products. In meiosis, homologous
chromosomes assort independently, resulting
in haploid cells with new combinations of
wild-type and mutant genes. By definition,
independent assortment holds for genes located on different chromosomes (different
linkage groups). However, the high frequency
of crossing-over that occurs during meiosis allows even closely linked genes to be separable.
If a large number of spores are scored (generated through either tetrad analysis or a random spore protocol), a strain with almost any
combination of markers can be isolated. Large
numbers of spores can be analyzed by random
spore analysis (Alternate Protocol).
The frequency with which linked markers
are separated by recombination is the basis for
genetic mapping in all organisms and is mainly
a function of the physical distance separating
them. This frequency is equal to twice the genetic distance in centimorgans (cM). Thus, two
genes separated by a genetic distance of 5 cM
will be recombined in 10% of all tetrads. In
yeast, a genetic distance of 1 cM corresponds
to a physical distance of ∼2.5 kb.
Critical Parameters
Diploids are usually selected by streaking
mating mixtures on minimal plates that are
supplemented (by spreading liquid stocks of
the nutrients on the surface of agar plates at the
concentrations listed in Table 13.1.1; see also
UNIT 1.3) with the known nutritional requirements predicted for the diploid. If diploids cannot be obtained by this method, make sure that
the haploid parents were of opposite mating
type. If they were, and the selective plates are
correct, the haploids probably carry at least one
additional nutrient requirement that is common to both. In this case, select on CM dropout
plates that select for two or more auxotrophic
requirements of the diploid (at least one auxotrophy for each haploid parent). It is impor-
tant that the haploid parents be from cultures
that have been freshly grown, but cells from
cultures stored 1 or 2 days at 4◦ C will also
mate well. If cells from plates that have been
stored longer are used, allow mating to proceed overnight.
In most laboratory strains of S. cerevisiae,
many diploid cells in a culture do not undergo
meiosis, resulting in a population of spores
contaminated to varying degrees with the original diploid. In addition, spores produced by
a single meiotic event are often notoriously
difficult to separate. The random spore protocol employs an extended incubation with
Zymolyase to destroy contaminating diploids,
followed by sonication and detergent treatment to disperse spores. The procedure should
yield a spore population with <1 diploid cell
per 104 spores. Spore colonies should be tested
for mating type (see Basic Protocol 5) to determine the frequency of diploid contamination (identified by their nonmating phenotype).
Contamination by diploids is usually not a
problem when tetrads are dissected, although
sometimes three spores and an unsporulated
diploid are accidentally dissected as a single
tetrad.
While most strains do not sporulate
synchronously, pregrowth in YPA medium
(UNIT 13.1) can result in a degree of synchrony
that is useful for monitoring gene expression
as meiosis progresses. For more precise synchrony, S. cerevisiae strain SKI can be used
(Esposito and Klapholz, 1981). This strain undergoes a highly synchronous meiosis which
it completes in 8 hr. For this reason, SKI and
its derivatives have been extremely useful for
studying the molecular and cellular events that
occur in meiosis (Wang et al., 1987).
When spores are allowed to grow up, it
is not uncommon to find tetrads where only
1, 2, or 3 of the spores have grown into
colonies. This should be relatively rare (<5%
of the tetrads) when closely related haploids
are used, but may occur at very high frequencies when the haploid parents are less closely
Yeast
13.2.11
Current Protocols in Molecular Biology
Supplement 82
related (as is often the case when strains from
different laboratories are mated). If one is simply trying to construct a useful genotype, this is
not a major problem. However, it is a problem
when viability of spore products is being analyzed to determine if a mutation is lethal.
If a marker is needed from a foreign strain
where spore viability is poor in crosses with
commonly used laboratory strains, spore products carrying that marker must be repeatedly crossed back to a laboratory standard
strain—the goal being to introduce only the desired mutation into the genetic background of
the laboratory strains. This procedure, called
backcrossing, is carried out until spore viability reaches acceptable levels (usually between
4 and 7 backcrosses). At each meiosis, spores
are identified that carry the foreign marker in
conjunction with several markers found in the
laboratory standard strain that is used as the
backcross parent. If the foreign marker has
been cloned, it is often more efficient to introduce a defined mutation into laboratory strains
by transformation, since each backcross
requires ∼10 days.
Time Considerations
Diploid selection will require ∼3 days.
Sporulation will take 2 to 4 days. Tetrad preparation will require ∼1 hr, and one should allot
several hours to learning how to dissect tetrads.
The dissected spores need ∼3 days of growth
before they form colonies that can be replica
plated for genotyping, which requires an additional overnight incubation on the appropriate
selective plates. Thus, between 9 and 11 days
are required to construct a new haploid strain
when starting from haploid parents. This time
does not include pregrowing diploids on YPD
medium prior to sporulation, which increases
sporulation efficiency.
Literature Cited
Esposito, R.E. and Klapholz, S. 1981. Meiosis and
ascosporal development. In The Molecular Biology of Yeast Saccharomyces: Life Cycle and
Inheritance ( J.N. Strathern, E.W. Jones, and J.R.
Broach, eds.) pp. 211-287. Cold Spring Harbor
Laboratory, Cold Spring Harbor, N.Y.
Sherman, F. 2002. Getting started with yeast. In
Guide to Yeast Genetics and Molecular and Cell
Biology, Part B (C. Guthrie and G.R. Fink, eds.)
pp. 3-41. Academic Press, San Diego.
Wang, H-T., Frackman, S., Kowalisyn, J., Esposito,
R.E., and Elder, R. 1987. Developmental regulation of SPO13, a gene required for separation
of homologous chromosomes at meiosis I. Mol.
Cell. Biol. 7:1425-1435.
Key Reference
Sherman, F., Fink, G.R., and Lawrence, C.W. 1979.
Methods in Yeast Genetics. Cold Spring Harbor
Laboratory, Cold Spring Harbor, N.Y.
Provides a number of detailed procedures for genetic experiments that may be of interest to more
advanced students.
Growth and
Manipulation of
Yeast
13.2.12
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Current Protocols in Molecular Biology
Genome-Wide Transposon Mutagenesis
in Yeast
UNIT 13.3
Transposon-based insertion screens provide a convenient means to identify gene function.
Transposon mutagenesis allows one to rapidly generate a large set of mutations that can
be screened for mutant phenotypes and specific patterns of gene expression. Moreover,
transposon-insertion alleles are much easier to detect than those generated by agents that
predominantly produce single-base changes. Insertional mutagenesis is most effectively
performed in yeast by means of shuttle mutagenesis. In this procedure, a library of yeast
genomic DNA is mutagenized with a bacterial transposon in vivo in E. coli; mutant alleles
are subsequently transferred into yeast for functional analysis. In yeast, shuttle mutagenesis is preferred over transposon mutagenesis in vivo, as prokaryotic transposons insert
more randomly than do endogenous yeast transposable elements.
This unit provides comprehensive protocols for the use of insertional libraries generated
by shuttle mutagenesis. From the Basic Protocol presented here, a small aliquot of
insertional library DNA may be used to mutagenize yeast, producing strains containing
a single transposon insertion within a transcribed and translated region of the genome.
This transposon-mutagenized bank of yeast strains may be screened for any desired
mutant phenotype. Alternatively, since the transposon contains a reporter gene lacking its
start codon and promoter, transposon-tagged strains may also be screened for specific
patterns of gene expression. Strains of interest may be characterized by vectorette PCR
(see Support Protocol 1) in order to locate the precise genomic site of transposon insertion
within each mutant. A method by which Cre/lox recombination may be used to reduce
the transposon in yeast to a small insertion element encoding an epitope tag is described
below (see Support Protocol 2). This tag serves as a tool by which transposon-mutagenized gene products may be analyzed further (e.g., localized to a discrete subcellular
site). For additional considerations regarding these transposon-mutagenized libraries (see
Strategic Planning).
NOTE: All solutions and equipment coming into contact with living cells must be sterile,
and aseptic technique should be used accordingly.
STRATEGIC PLANNING
At present, the authors have generated three different transposon-mutagenized libraries
of yeast genomic DNA. Each library presents its own unique advantage as an insertional
mutagen; therefore, care should be taken in choosing an appropriate library when planning
a genome-wide screen for gene function.
Specifically, two mutagenized libraries contain transposon insertions bearing the reporter
gene, lacZ, lacking both its promoter and initiator ATG codon. Our first lacZ-insertional
library (Burns et al., 1994) was derived by transposon mutagenesis of a genomic library
constructed in the Tn3-free vector pHSS6 (Seifert et al., 1986); each library plasmid
contains a 3 to 4 kb fragment from a Sau3A partial digest of yeast genomic DNA. This
library (containing 18 genome equivalents of DNA) was mutagenized in vivo in E. coli
with mTn-lacZ/LEU2, a Tn3-derived minitransposon (mTn) carrying the yeast selectable
marker LEU2 and a bacterial marker encoding resistance to ampicillin (Fig. 13.3.1). The
resulting insertional library exhibits a random distribution of transposon-tagged genes,
but does contain 2µm plasmid DNA.
Saccharomyces
cerevisiae
Contributed by Anuj Kumar and Michael Snyder
Current Protocols in Molecular Biology (2000) 13.3.1-13.3.15
Copyright © 2000 by John Wiley & Sons, Inc.
13.3.1
Supplement 51
A
TR
lacZ
LEU2
amp
TR
• ampicillin resistance (ampr)
• LEU2 selection
mTn-lacZ/LEU2
• IacZ reporter lacking a
promoter and ATG codon
B
loxR
TR
lacZ
(tet r)
URA3
loxP
TR
3xHA
mTn-3xHAllacZ
Cre-lox
recombination
• tetracycline resistance (tetr)
• URA3 selection
• IacZ reporter lacking a
promoter and ATG codon
274-bp
(93-AA HAT tag)
• HAT-tagging by Cre-lox
recombination
C
loxR
TR
GFP
URA3
(tet r)
loxP
TR
3xHA
mTn-3xHA/GFP
• tetracycline resistance
Cre-lox
recombination
• URA3 selection
• full-length GFP reporter
274-bp
(93-AA HAT tag)
Genome-Wide
Transposon
Mutagenesis in
Yeast
• HAT-tagging by Cre-lox
recombination
Figure 13.3.1 Transposons for genome-wide random mutagenesis in yeast. Each indicated
transposon has been used to mutagenize a plasmid-based library of yeast genomic DNA in E. coli.
Relevant features of each transposon are indicated below the corresponding schematic. (A)
mTn-lacZ/LEU2 Tn3-derived minitransposon capable of generating disruption alleles and reporter
gene fusions. Proteins tagged by mTn-lacZ/LEU2 insertion may be immunolocalized using antibodies directed against β-gal. (B) mTn-3xHA/lacZ. Cre/lox recombination may be used to reduce
mTn-3xHA/lacZ in yeast to a 274-bp epitope-insertion element. Due to a 5-bp duplication in
target-site sequence associated with Tn3 transposition, this insertion element encodes a 93-amino
acid tag containing three copies of the HA epitope. mTn-3xHA/lacZ may be used to generate
lacZ-fusions, disruption alleles, conditional alleles, and HA-tagged alleles. (C) mTn-3xHA/GFP, a
minitransposon carrying full-length GFP reporter gene, in other respects, identical to mTn3xHA/lacZ. Note both mTn-3xHA/GFP and mTn-3xHA/lacZ encode identical HA tags upon Cre/lox
recombination. Corresponding insertion libraries may be requested at the authors’ Web site (see
Internet Resources).
13.3.2
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Current Protocols in Molecular Biology
A second lacZ-insertional library was generated by mutagenesis with a URA3-marked
mTn3 transposon (mTn-3xHA/lacZ; Ross-Macdonald et al., 1997) modified to incorporate lox elements and sequence encoding three copies of an epitope from the influenza
virus hemagglutinin protein (the HA epitope; Fig. 13.3.1). Cre-mediated recombination
between lox sites (see Support Protocol 2) may be used to reduce the 6 kb mTn-3xHA/lacZ
insertion to a smaller element encoding an in-frame epitope tag; this allele may be used
to determine the subcellular localization of transposon-tagged gene products. This small
93-amino acid epitope tag is also an effective means of generating conditional alleles and
hypomorphic mutants for further study. Using mTn-3xHA/lacZ, we have mutagenized a
plasmid library (containing 50 genome equivalents) generated by Sau3A partial digestion
of genomic DNA isolated from a cir0rho0 yeast strain. The resulting mTn-3xHA/lacZ
insertion library, therefore, lacks 2µ plasmid sequences and mitochondrial-encoded DNA.
This mTn-3xHA/lacZ insertional library has been used in the bulk of our genomic studies
with excellent results (Ross-Macdonald et al., 1999); however, it does possess a less
random distribution of transposon insertions as compared to that found in the mTnlacZ/LEU2 library.
A third insertion library was generated by mutagenesis with a mini-Tn3 transposon
(mTn-3xHA/GFP; Ross-Macdonald et al., 1997) encoding full-length green fluorescent
protein (GFP) in place of β-galactosidase (Fig. 13.3.1); in all other respects, this
transposon is identical to mTn-3xHA/lacZ. GFP is a versatile reporter facilitating a variety
of biological studies in living cells (e.g., fluorescence microscopy, fluorescence-flow
cytometry, and fluorescent tagging as a means of determining protein localization). We,
however, have found lacZ to be a more sensitive reporter of gene expression and, therefore,
more effective in analyzing genes expressed at a low level.
Diagrams of each transposon, with accompanying summaries, are presented in Figure
13.3.1.
Several considerations should be addressed in choosing a yeast host strain. If using an
insertional library derived from mutagenesis with mTn-lacZ/LEU2, choose a Leu2− yeast
strain. Use of a diploid strain will allow recovery of transposon insertions within essential
genes. Also, the use of a cir0 strain will prevent recovery of insertions within the yeast 2µ
plasmid (applicable to the mTn-lacZ/LEU2- and mTn-3xHA/GFP-mutagenized libraries,
which were not generated from a cir0 strain). For the eventual analysis of HAT-tagged
proteins (Support Protocol 2), choose a Ura3−, Leu2− strain of yeast; preferably transform
this strain with pGAL-cre (pB227; see Support Protocol 2) prior to use in this protocol.
Irrespective of the host strain chosen, yeast cultures should be grown to mid-log phase in
order to ensure maximal transformation efficiency.
Saccharomyces
cerevisiae
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Current Protocols in Molecular Biology
Supplement 51
BASIC
PROTOCOL
GENERATING YEAST MUTANTS FROM mTn-MUTAGENIZED
LIBRARY DNA
This protocol describes methods by which a transposon-mutagenized library of yeast
genomic DNA may be used to produce a bank of yeast strains, each strain carrying a
transposon insertion within protein-coding sequence. Specifically, the following protocol
assumes use of the mTn-3xHA/lacZ-mutagenized library (see Strategic Planning), although an identical approach could be applied to the manipulation of any lacZ-based
insertional library.
The mTn-3xHA/lacZ/URA3-derived insertional library (as well as any other library
described above) may be requested using order forms linked to the authors’ Web page
(http://ygac.med.yale.edu; see Internet Resources). Upon receipt of this library, mutagenized DNA may be excised from pHSS6 for transformation into an appropriate Ura3−
yeast strain according to protocols modified from Chen et al. (1992). By homologous
recombination, the mTn-mutagenized genomic DNA fragment will replace its chromosomal locus, generating a yeast strain carrying a chromosomal mTn insertion. Transformants carrying an in-frame fusion of mTn-encoded lacZ to yeast protein coding sequence
may be identified using a color assay for β-galactosidase (β-gal) activity. The resulting
bank of yeast strains containing productive lacZ fusions may then be screened for any
desired mutant phenotype.
Materials
Transposon-mutagenized genomic library plasmid DNA (available upon request;
see Internet Resources)
10× TE buffer, pH 8.0 (APPENDIX 2), sterile
E. coli tets, kans (e.g., DH5α)
14-cm LB plates and medium supplemented with 3 µg/ml tetracycline and 40
µg/ml kanamycin (UNIT 1.1)
LB medium (UNIT 1.1)
Glycerol, sterile
NotI restriction endonuclease and buffer (UNIT 3.1)
Ura3− yeast culture (Chapter 13)
One-step buffer (see recipe)
10 mg/ml denatured salmon sperm DNA (UNIT 14.7)
CM dropout plates and medium without uracil (–Ura; UNIT 13.1)
YPAD plates: YPD plates (UNIT 13.1) supplemented with 80 mg/liter adenine
Chloroform
Xgal plates (UNIT 13.1)
Clinical tabletop centrifuge
45°C water bath or incubator
Sterile toothpicks
3MM filter paper (Whatman)
30°C incubator
9-cm and 15-cm glass petri dishes
Genome-Wide
Transposon
Mutagenesis in
Yeast
Additional reagents and equipment for culturing E. coli (UNITS 1.1, 1.2 & 1.3),
preparing and transforming competent E. coli (UNIT 1.8), isolating plasmid DNA
by miniprep (UNIT 1.6) or large-scale preparation (UNIT 1.7), restriction
endonuclease digestion (UNIT 3.1), yeast culture (UNIT 13.2), agarose gel
electrophoresis (UNIT 2.5), culturing yeast (UNITS 13.1 & 13.2), transforming yeast
(UNIT 13.7), and assaying for β-galactosidase activity (UNIT 13.6)
13.3.4
Supplement 51
Current Protocols in Molecular Biology
Prepare library DNA
1. Distribute plasmid DNA from individual pools of the transposon-mutagenized
genomic library (∼1 µg DNA per pool) as dried-down solutions. Briefly centrifuge
each dry sample, and resuspend DNA from each pool in an appropriate volume of
TE buffer, pH 8.0 (e.g., 10 µl per pool). Each pool of library DNA is mutagenized
independently; therefore, it is advisable to process DNA from each pool separately
to ensure independent insertion alleles are obtained.
2. Introduce a suitable amount of DNA from each pool into any tetracycline– (tets) and
kanamycin– sensitive (kans) E. coli strain by standard transformation procedures (UNIT
1.8).
Standard E. coli strains (e.g., DH5α) are suitable hosts for this transformation. Electroporation (UNIT 9.3) is not required, although resulting transformant yields may be increased
10- to 100-fold. If electroporation is used, decrease DNA quantity appropriately.
3. Select transformants on LB plates (14 cm in diameter) supplemented with 3 µg/ml
tetracycline and 40 µg/ml kanamycin.
Approximately 10,000 transformants should be obtained per pool following overnight
growth at 37°C.
4. Elute transformant colonies by placing 6 ml of LB medium onto the surface of each
plate and scraping cells into a homogenous suspension.
An aliquot of this suspension should be stored at −70°C in 15% glycerol.
5. Dilute a 1-ml aliquot of this eluate into 100 ml LB medium supplemented with 3
µg/ml tetracycline and 40 µg/ml kanamycin to yield a culture of nearly saturated cell
density. Incubate at 37°C with aeration for 2 to 3 hr.
The remaining eluate may be stored at 4°C until completion of subsequent steps.
6. Isolate plasmid DNA by standard miniprep (UNIT 1.6) or large-scale preparation (UNIT
1.7).
Transform yeast with mTn-mutagenized library DNA
7. Digest a small aliquot (typically, at least 1 µg) of plasmid DNA from each library
pool (obtained in step 6) with NotI restriction endonuclease (UNIT 3.1). Subsequently,
analyze a portion of the reaction mixture by agarose gel electrophoresis (UNIT 2.5) to
ensure release of mTn-mutagenized yeast DNA from the pHSS6 vector. Store the
remaining reaction mixture at 4°C for later use (step 10).
Upon electrophoresis, a distinct 2.1-kb band (pHSS6) and broad 8-kb band (mTn-mutagenized yeast genomic DNA) should be visible.
The broad 8-kb band generated by NotI digestion consists of 2- to 4-kb inserts of yeast
genomic DNA carrying the 6-kb transposon construct. As this library was constructed from
size-fractionated yeast genomic DNA fragments, insert sizes are relatively homogeneous.
8. Grow a 10-ml culture of any desired Ura3− yeast strain to mid-log phase (a density
of 107 cells/ml or OD600 of ∼1) maintaining appropriate selection, as applicable.
Several considerations should be addressed in choosing a yeast host strain (see Strategic
Planning).
9. Centrifuge cells in a clinical tabletop centrifuge for 5 min at 1100 × g at room
temperature. Wash pellet once with 5 vol of one-step buffer.
This initial wash in one-step buffer is particularly important if culture volumes are
increased.
Saccharomyces
cerevisiae
13.3.5
Current Protocols in Molecular Biology
Supplement 51
10. Resuspend cells in 1 ml of one-step buffer supplemented with 1 mg of denatured
salmon sperm DNA. Add 100-µl aliquots from this suspension to microcentrifuge
tubes containing 0.1 to 1 µg of NotI-digested plasmid DNA from step 7. Vortex tubes
to mix contents thoroughly.
To minimize generation of transformants containing more than one insertion, use just
enough transforming DNA to yield a reasonable number of transformants (see Anticipated
Results). For best results, perform this pilot experiment after determining optimal conditions; scale up as appropriate.
11. Incubate this mixture at 45°C for 30 min.
12. Microcentrifuge cells for 5 sec at maximum speed, room temperature, and subsequently resuspend pellet in 400 µl of CM (–Ura) dropout medium. Spread 200-µl
aliquots onto CM dropout plates (−Ura). Incubate at 30°C for 3 to 4 days.
Up to 1 × 103 transformants may be recovered per 1 ìg of transforming DNA. If the
mTn-lacZ/LEU2-mutagenized library and leu2 host strain are used, recover transformants
on appropriate CM without leucine (−Leu) dropout medium.
Screen transformants for β-galactosidase activity
13. To maximize detection of lacZ-fusions expressed at a low level, transfer transformant
colonies onto YPAD plates with sterile toothpicks (UNIT 13.2) at a density of up to 100
colonies per plate.
If an Ade2− host strain is used, the addition of adenine to growth medium will prevent any
accumulation of red pigment; otherwise, accumulated red pigment may obscure the blue
color produced in subsequent assays for β-galactosidase (β-gal) activity.
14. Place a sterile disc of 3MM filter paper onto a CM dropout plate –Ura; repeat for as
many plates as desired. Replicate transformant cells onto filter-covered plates, to
allow easy identification of corresponding colonies on YDAP plates (step 13), and
incubate overnight at 30°C.
As some transposon insertions may impair growth even in the heterozygous state, selection
for uracil prototrophy should be maintained during this period of vegetative growth.
Similarly, selection for leucine prototrophy should be maintained if using a Leu2− host.
Other growth conditions (e.g., growth on sporulation medium) may be substituted above
as desired.
15. Following overnight growth, lift filters from plates and place in the lid of a 9-cm glass
petri dish. Place this lid inside a closed 15-cm glass petri dish containing chloroform.
Incubate for 10 to 30 min.
The minimum exposure time required to lyse a particular yeast strain must be determined
empirically.
16. Place filters colony-side-up onto Xgal plates. Incubate inverted at 30°C for up to 2
days.
To minimize cost, very thin plates containing Xgal may be used with no loss in signal quality.
The signal intensity observed using an Xgal plate is superior to that obtained by soaking
the filters in a buffered Xgal solution.
17. Recover transformants producing β-gal (indicated by blue staining on Xgal plates)
from the regrown YPAD plates generated in step 13. Maintain appropriate selection
during subsequent growth and manipulation of strains.
Genome-Wide
Transposon
Mutagenesis in
Yeast
If desired, strains of interest may be stored long-term in 15% glycerol at −70°C following
overnight growth at 30°C in appropriate CM dropout medium.
13.3.6
Supplement 51
Current Protocols in Molecular Biology
VECTORETTE POLYMERASE CHAIN REACTION
Banks of transposon-mutagenized yeast strains (see Basic Protocol) represent powerful
tools by which genetic screens can be performed rapidly on a genome-wide scale. Large
numbers of yeast mutants exhibiting a given disruption phenotype may be easily identified
in these genome-wide screens; however, in order for this analysis to be informative, the
precise genomic site of mTn insertion must be located within each of these transformants.
Accordingly, the following support protocol provides an adaptation of the vectorette
polymerase chain reaction (PCR) method of Riley et al. (1990) by which genomic DNA
from any site of mTn insertion may be easily recovered for subsequent DNA sequence
analysis.
SUPPORT
PROTOCOL 1
In vectorette PCR, genomic DNA is initially digested with any suitable blunt-end
restriction endonuclease possessing a 4- to 6-base-pair recognition sequence. Following
digestion, DNA fragments are ligated to a pair of annealed primers containing a non-homologous central region; these primer pairs form “anchor bubbles” flanking each genomic
fragment. PCR is then performed using a primer complementary to mTn sequence and a
primer identical to sequence within the anchor bubble. During the initial round of
amplification, only the mTn primer can bind its template; however, during subsequent
cycles, the anchor bubble primer can anneal to the elongated mTn primer, resulting in
selective amplification of DNA sequence adjacent to the point of mTn insertion. Vectorette
PCR is summarized in Figure 13.3.2.
Materials
Anchor bubble primers 1 and 2 (see Fig. 13.3.2 for sequences; see UNITS 2.11 & 2.12
for synthesis techniques)
1 M MgCl2 (APPENDIX 2)
Universal vectorette (UV) and mTn primers (see Fig. 13.3.2 for sequences; see
UNITS 2.11 & 2.12 for synthesis techniques)
Transposon-mutagenized yeast strain (see Basic Protocol)
Appropriate restriction endonucleases (e.g., AluI or DraI) and buffers (UNIT 3.1)
10× T4 DNA ligase buffer (UNIT 3.4)
5 mM ATP (UNIT 3.4)
400 U/µl T4 DNA ligase (measured in cohesive end ligation units)
5 U/µl Taq DNA polymerase and 10× buffer (UNITS 3.5 & 15.1)
2.5 mM 4dNTP mix (UNIT 15.1)
Heat block
Automated thermal cycler
Additional reagents and equipment for preparing yeast DNA (UNIT 13.11), restriction
endonuclease digestion (UNIT 3.1), ligation of DNA fragments (UNIT 3.16), PCR
amplification (UNIT 15.1), purification of DNA from agarose gels (UNIT 2.6), and
DNA sequence analysis (UNIT 7.4)
1. In preparation for vectorette PCR, synthesize anchor bubble primers (suggested DNA
sequence indicated in Fig. 13.3.2). Form anchor bubble as follows.
a. Prepare an aqueous solution that contains 2 to 4 mM each of anchor bubble primers
1 and 2.
b. Denature by incubating 5 min at 95°C in a heat block.
c. Add 1 M MgCl2 to a final concentration of 2 mM.
d. Anneal by removing the block from the base unit and placing it on the bench top
until it slowly cools to room temperature.
The annealed anchor bubble may be stored at −20°C until step 5.
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cerevisiae
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Current Protocols in Molecular Biology
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2. Synthesize universal vectorette (UV) primer (suggested DNA sequence indicated in
Fig. 13.3.2). Synthesize primer complementary to the mTn used to generate the
insertion library (mTn primer).
If using a lacZ-insertional library, the following sequence is recommended for the mTn
primer: 5′-CGC CAG GGT TTT CCC AGT CAC GAC-3′.
The UV primer is identical in sequence (not complementary) to a portion of anchor bubble
primer 2 (Fig. 13.3.2). The mTn primer indicated above corresponds to sequence from the
5′-end of lacZ on the antisense strand of mTn-3xHA/lacZ. If analyzing a mTn-3xHA/GFPmutagenized strain, the following GFP primer sequence is recommended: 5′-CAT CAC
CTT CAC CCT CTC CAC TGA C-3′.
A
mTn3
Digest DNA
Ligate bubble
Yeast st rain
with mTn
insertion
mTn3
PCR: cycle 1
M13 primer
binds template
mTn
PCR: subsequent
cycles
UV
mTn
UV primer binds extended mTn product;
amplification of region between
UV and M13 primers
B
Anchor bubble primer-1: GAAGGAGAGGACGCTGTCTGTCGAAGGTAAGGAACGGACGAGAGAAGGGAGAG
Anchor bubble primer-2: GACTCTCCCTTCTCGAATCGTAACCGTTCGTACGAGAATCGCTGTCCTCTCCTTC
UV primer: CGAATCGTAACCGTTCGTACGAGAATCGCT
mTn primer: CGCCAGGGTTTTCCCAGTCACGAC
Genome-Wide
Transposon
Mutagenesis in
Yeast
Figure 13.3.2 Vectorette PCR. (A) In preparation for vectorette PCR, small, blunt-ended fragments of mTn-mutagenized genomic DNA are generated by restriction enzyme digestion; DNA
fragments are subsequently ligated to preannealed anchor bubble primers. Amplification is carried
out using mTn and Universal vectorette (UV) primers (sequence indicated). During the first cycle
of PCR, only the mTn primer can bind to template, as the UV primer is identical (not complementary)
to anchor bubble sequence. During subsequent PCR cycles, the UV primer can bind to elongated
product from the mTn primer, resulting in the selective amplification of genomic DNA adjacent to
the mTn insertion site. Primers and major PCR products are shown in bold. (B) Suggested anchor
bubble and PCR primers are listed below: sequence is indicated 5′ to 3′. The Universal vectorette
(UV) primer is identical to the boxed sequence within Anchor Bubble primer-2.
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Current Protocols in Molecular Biology
3. Prepare genomic DNA from a transposon-mutagenized yeast strain of interest (UNIT
13.11).
Care should be taken to obtain high-quality DNA; this is critical for successful PCR
amplification.
4. Digest 1 to 3 µg of genomic DNA overnight at 37°C with 10 U of either AluI or DraI
in a total reaction volume of 20 µl (UNIT 3.1). Following digestion, incubate at 65°C
for 20 min to inactivate the enzyme.
Ideally, genomic DNA fragments should be small and blunt-ended in preparation for
subsequent anchor bubble ligation; accordingly, any restriction enzyme used in step 4
should cut frequently and leave blunt-ends after cleavage.
IMPORTANT NOTE: The enzyme must not cut between the transposon end and the mTn
primer binding site.
5. To the reaction mix from step 4, add the following:
5 µl 10× T4 DNA ligase buffer
22.5 µl sterile water
1 µl annealed anchor bubble (generated in step 1)
0.5 µl 5 mM ATP
1 µl (400 U) T4 DNA ligase.
Incubate 9 to 24 hr at 16°C.
To accommodate blunt-end ligation, incubation times have been extended up to 24 hr as
indicated. The addition of polyethylene glycol to a final concentration of 15% may enhance
blunt-end ligation.
6. Prepare PCR mix (100 µl total) as follows.
a. Withdraw 5 µl from the ligation reaction from step 4.
b. To this 5-µl aliquot, add:
10 µl 10× Taq PCR buffer
71 µl sterile water
8 µl 2.5 mM dNTP mix
2.5 µl 20 µM mTn primer (step 2)
2.5 µl 20 µM UV primer (step 2)
1 µl (5 U) Taq DNA polymerase.
If desired, the mixture above may be modified for “hot start” PCR (UNIT 15.1) using wax
beads commercially available from a variety of sources; please follow manufacturer-suggested guidelines.
7. Transfer to an automated thermal cycler and denature at 92°C for 2 min. Carry out
PCR (UNIT 15.1) under the following conditions:
35 cycles:
1 cycle:
20 sec
30 sec
45 sec
90 sec
92°C
67°C
72°C
72°C
(denaturation)
(annealing)
(extension)
(extension)
In our experience, short cycling times tend to improve product yield; however, optimum
conditions for PCR amplification must be determined empirically.
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cerevisiae
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Current Protocols in Molecular Biology
Supplement 51
8. Analyze 80 µl of the reaction mix from step 7 by nondenaturing agarose gel
electrophoresis (UNIT 2.7); a single band containing 200 to 400 ng of DNA should be
visible. Excise the PCR product and recover DNA in TE buffer (∼12 µl) as described
in UNIT 2.6.
If multiple bands are present, excise all bands and recover DNA individually from each.
DNA may be recovered with Qiaex resin (available from Qiagen) if desired.
9. Analyze ∼4 to 6 µl of recovered product by DNA sequencing (UNIT 7.4) to identify the
exact site of mTn insertion.
SUPPORT
PROTOCOL 2
EPITOPE TAGGING OF mTn-MUTAGENIZED GENE PRODUCTS
Transposon-mutagenized yeast strains may be analyzed further by utilizing the epitopetagging feature incorporated into mTn-3xHA/lacZ and mTn-3xHA/GFP. As shown in
Figure 13.3.1, these minitransposons each contain a pair of lox elements located internal
to the sequence, encoding three copies of the HA epitope. These lox sequences are target
sites for the Cre recombinase, which catalyzes site-specific recombination between the
lox sites. Therefore, expression of Cre results in excision of the central body of the
transposon, leaving behind a 274-bp sequence containing the HA-epitope coding region.
Due to a 5-bp target site duplication associated with Tn3 transposition, this reduced
construct corresponds to a small element encoding 93 amino acids (the HA-epitope tag
or HAT tag) inserted within the protein. This HAT tag insertion element is an effective
means of generating conditional alleles, hypomorphic mutants, and epitope-tagged strains
for immunodetection.
The following support protocol describes a method by which yeast strains bearing an
in-frame mTn insertion may be used to derive corresponding HAT-tagged strains by
Cre/loxP recombination in vivo. The phage P1 Cre recombinase is expressed exogenously
from plasmid pGAL-cre (available from authors’ Web site; see Internet Resources). On
this plasmid, cre is under transcriptional control of the GAL promoter; therefore, induction
by galactose can be used to drive cre expression. Following galactose induction, cells that
have undergone Cre-mediated recombination (and loss of the URA3 marker) are selected
on medium containing 5-fluoroorotic acid (5-FOA). In the authors’ experience, galactose
induction has resulted in Cre-mediated excision of the mTn-encoded URA3 marker in
>90% of cells analyzed.
Materials
mTn-mutagenized yeast strain (see Basic Protocol)
pGAL-cre (pB227; available upon request from authors, see Internet Resources)
Raff/CM dropout plates and medium –Leu, –Ura with 2% (w/v) raffinose (UNIT 13.1)
Gal/CM dropout medium –Leu with 2% (w/v) galactose (UNIT 13.1)
Glc/CM dropout medium –Leu with 2% (w/v) glucose (UNIT 13.1)
5-fluoroorotic acid plates (5-FOA; UNIT 13.1)
Glycerol, sterile
30°C shaker and incubator
Additional reagents and equipment for culture of yeast (UNIT 13.2) and
transformation of yeast cells (UNIT 13.7)
Genome-Wide
Transposon
Mutagenesis in
Yeast
13.3.10
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Current Protocols in Molecular Biology
1. Transform mTn-mutagenized yeast strain with pGAL-cre (pB227) using standard
procedure (see Basic Protocol or UNIT 13.7). Select transformants on CM –Leu, –Ura
dropout plates.
Alternatively, pGAL-cre may be transformed into a desired yeast strain prior to transformation with mTn-mutagenized genomic DNA (see Basic Protocol, steps 8 to 11).
This plasmid contains the following elements: amp, ori, CEN, and LEU2.
2. Inoculate transformants into 2 ml Raff/CM dropout medium –Leu, –Ura with 2%
raffinose as the carbon source. Incubate at 30°C with aeration until the culture has
grown to saturation.
Growth in raffinose derepresses the GAL promoter.
3. Dilute cultures 100-fold into 2 ml Gal/CM –Leu dropout medium with 2% galactose
as the carbon source. As a control, dilute an aliquot of this same culture 100-fold into
2 ml Glc/CM –Leu, with 2% glucose as the carbon source. Grow cultures for 2 days
at 30°C with aeration.
Some strains grow very poorly in galactose; however, galactose induction is sometimes
effective even without visible signs of growth.
4. Test strains for loss of URA3 marker as follows.
a. If visible growth is apparent in cultures grown on 2% galactose, dilute cultures
100-fold in sterile water and withdraw a 10-µl aliquot.
b. If no growth is apparent in cultures grown on 2% galactose, withdraw a 10-µl
aliquot from the undiluted culture.
c. Dilute cultures grown in 2% glucose 100-fold in sterile water and withdraw a 10-µl
aliquot.
Spot the aliquot onto a 5-FOA plate; isolate single colonies by streaking the droplet.
Incubate 5-FOA plates at 30°C until growth is visible on those plates inoculated with
strains grown in galactose.
Loss of the transposon-encoded URA3 gene by galactose induction of the Cre recombinase
should be reflected in colony growth on 5-FOA plates containing strains grown in
galactose. 5-FOA plates carrying strains grown in the presence of glucose should display
little or no growth, as expression of the Cre recombinase is repressed by glucose.
Alternatively, plate diluted cultures obtained in this step onto CM medium and replicate
on CM dropout medium –Ura. Incubate at 30°C ∼2 days. Cultures grown in galactose
should yield roughly 100-fold more Ura– cells than identical cultures grown in glucose.
5. Save single colonies from strains that have lost the URA3 (marker exclusively
following galactose induction) as a stock in 15% (w/v) glycerol at −70°C.
If desired, PCR analysis may be used to confirm the position of the HAT tag; the complete
274-bp DNA sequence encoding this tag may be viewed on the authors’ Web site (see
Internet Resources).
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cerevisiae
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Current Protocols in Molecular Biology
Supplement 51
REAGENTS AND SOLUTIONS
Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see
APPENDIX 2; for suppliers, see APPENDIX 4.
One-step buffer
0.2 M lithium acetate
40% (w/v) PEG 4000
100 mM 2-mercaptoethanol
Store up to 1 week at 4°C in a dark bottle
COMMENTARY
Background Information
Genome-Wide
Transposon
Mutagenesis in
Yeast
Transposable elements are powerful insertional mutagens capable of efficiently generating large numbers of informative mutant alleles
for subsequent functional analysis. In particular, transposon-based mutagenesis systems offer several advantages over conventional mutagenesis techniques discussed in UNIT 13.3 (i.e.,
treatment with ethylmethane sulfonate and UV
irradiation). In contrast to these approaches,
transposon mutagenesis may be used to generate a diverse variety of insertion alleles within
target DNA; transposons may be modified to
incorporate reporter genes, regulatory elements, and epitope-tagging elements generating null alleles, reporter fusions, mis-expression alleles, conditional alleles, and epitopetagged alleles. Transposon insertions are
additionally advantageous in that they tag mutant loci with an easily detectable DNA element, thereby facilitating subsequent identification and analysis of mutated genes.
Several transposon systems are applicable
to genome-wide mutagenesis in yeast. Endogenous yeast Ty-based transposable elements
may be used for in vivo as well as in vitro
transposon mutagenesis (Garfinkel and Strathern, 1991; Devine and Boeke, 1994); however,
the utility of these systems may be limited by
strong target bias associated with Ty transposition in vivo (Ji et al., 1993) and poor transposition efficiency in vitro. Prokaryotic
transposons, such as Tn3 and Tn7, generally
exhibit less bias in target-site selection and offer
high mutation frequencies. Additionally, negative target regulation of bacterial transposition
renders DNA molecules already containing a
sin gle transposon immune to further
transposon insertion. Collectively, these properties of bacterial transposons enable them to
generate large numbers of single-hit mutants—
a suitable population for further functional
analysis.
Using shuttle mutagenesis, yeast DNA may
be mutagenized in E. coli with bacterial
transposons; transposon-mutagenized DNA
may then be returned to yeast by transformation
and homologous recombination (Hoekstra et
al., 1991). This unit describes the application
of Tn3-based shuttle mutagenesis as a means
of generating a bank of transposon-mutagenized yeast strains for functional analysis
in yeast. The Basic Protocol provides methods
by which a Tn3-mutagenized genomic DNA
library may be used to transform yeast, generating a bank of marked genomic mutations.
This mutant bank may be subsequently
screened for desired disruption phenotypes and
sp ecific p atterns of gene expression.
Transposon insertion sites may be identified in
strains of interest either by vectorette PCR
(Support Protocol 1) or by plasmid rescue
(Burns et al., 1994). Plasmid rescue protocols
may be found at the authors’ web site
(http://ygac.med.yale.edu; see Internet Resources). Cre/lox recombination may be induced within yeast strains mutagenized by
mTn-3xHA/lacZ or mTn-3xHA/GFP (Support
Protocol 2); resulting epitope-tagged gene
products may be immunolocalized with antibodies directed against the transposon-encoded
epitope. Epitope-tagging is also an effective
means of generating hypomorphic mutants exhibiting partial gene function.
These transposon insertion libraries constitute a valuable laboratory reagent facilitating
genetic screening in yeast. Traditionally, genetic studies have followed a paradigm in
which labor-intensive procedures such as complementation analysis or genetic mapping have
been necessary as a means of identifying the
affected gene within a mutant of interest.
Transposon-mutagenized strain collections, as
described in this unit, provide a means of altering this paradigm. Researchers may now identify affected genes within clones of interest
13.3.12
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Current Protocols in Molecular Biology
either by directly screening a bank of defined
mutants or by simply identifying the mutated
genomic loci; in either case, the mutated gene
can be identified rapidly with a minimal investment in time and effort. Such labor-saving
methodologies are essential in order to expedite
studies of gene function.
Critical Parameters and
Troubleshooting
Transposon-insertion libraries may be used
as insertional mutagens in yeast through a series of simple and straightforward steps. Insertion library DNA may be amplified in E. coli
using any standard method of DNA transformation (e.g., “heat-shock” treatment, electroporation). Transformants should be recovered
on growth medium supplemented with antibiotics as indicated; note that tetracycline should
be used at a final concentration of 3 µg/ml.
Mutagenized DNA from the library is excised
from its vector and subsequently transformed
into an appropriate yeast strain; electroporation
is not normally required, although its use can
increase transformant yields 10- to 100-fold.
Transform yeast with a small quantity of DNA
in order to minimize recovery of double integrants. Diploid strains containing multiple
transposon insertions may be identified by examining segregation of the transposon-encoded selectable marker (i.e., URA3 or LEU2)
upon tetrad dissection. Alternatively, Southern
analysis (UNIT 2.9) may be used to identify strains
containing two different transposon insertions.
For purposes of this analysis, lacZ-bearing
transposons may be probed with a 817-bp
BamHI-ClaI fragment from the 5′ end of lacZ.
Care should be taken to ensure that mutant
phenotypes of interest are linked to the
transposon insertion, as multiple insertion
events or unrelated spontaneous mutations may
have occurred during yeast transformation.
Within strains of interest, genetic analysis
should be used to confirm segregation of the
desired phenotype with the transposon-encoded marker.
The vectorette PCR (see Support Protocol
1) described in this unit allows efficient amplification of genomic DNA at the site of mTn
insertion. Typical of any PCR reaction, product
yield is dependent upon the quality of DNA
used as template. High-quality genomic DNA
may be obtained by any standard protocol;
CsCl purification of prepared DNA is not required. To function as template for vectorette
PCR, genomic DNA fragments must be modi-
fied by anchor bubble ligation. To facilitate
blunt-ended ligation, polyethylene glycol may
be added to the ligation mixture at a final
concentration of 15% (w/v). Longer ligation
times (in excess of 16 hr) may be beneficial.
Insufficient product yield may also be addressed by trial-and-error modification of indicated cycling conditions. Per cycle, primer extension at 72°C may be increased from 45 sec
to 3 min, if no PCR product is apparent; however, the authors typically have obtained better
results using shorter extension times. Alternatively, annealing conditions (suggested temperature of 67°C per cycle) may be made more
or less stringent as needed. If multiple PCR
products are present after amplification, each
product may be individually analyzed by DNA
sequencing.
Anticipated Results
Amplification of library DNA in E. coli
should generate roughly 10,000 colonies per
pool. Subsequent transformation of library
DNA into yeast should yield at least 2 × 105
transformants. Roughly 180,000 yeast transformants should be screened for reporter activity in order to ensure 95% coverage of the
genome; the authors typically observe β-gal
activity in 12% to 16% of transformants. For
purposes of insertional mutagenesis without
regard to in-frame reporter activity, 30,000 to
50,000 colonies should be screened
Within yeast transformants of interest,
transposon insertion sites may be identified by
vectorette PCR (Support Protocol 1). Vectorette
PCR can be expected to yield 200 to 400 ng of
product. This quantity of DNA should constitute sufficient template for 1 or 2 sequencing
reactions.
Transposon-mutagenized gene products
may be HAT-tagged through galactose-induced
Cre/lox recombination. The authors have generally observed poor growth of yeast strains on
medium containing galactose as its carbon
source; however, after growth for a few generations in galactose-containing medium, greater
than 90% of yeast cells exhibit loss of the mTn
URA3 marker. In contrast, less than 1% of cells
grown in the presence of glucose undergo recombination between mTn-encoded lox sites.
From a pilot study of epitope-tagged proteins,
the authors estimate that 40% to 75% of HATtagged proteins should be fully functional,
while ∼90% should localize properly within the
cell (Ross-Macdonald et al., 1997).
Saccharomyces
cerevisiae
13.3.13
Current Protocols in Molecular Biology
Supplement 51
Time Considerations
The protocols presented herein are neither
time-consuming nor labor-intensive. Large
quantities of library DNA may be prepared as
described within a period of 2 days. Library
DNA may be subsequently digested and transformed into yeast in one day; allow 3 to 4 days
for growth of transformants. Yeast mutants may
be incubated under desired growth conditions
for a length of time appropriate to the given
study (e.g., overnight). Subsequently, strains
may be assayed for β-gal activity in <1 hr; blue
staining may develop over a period of up to 2
days.
Transposon-mutagenized gene products
may be epitope-tagged through Cre/lox recombination over a period of ∼5 days; this protocol,
however, requires very little “hands-on” time.
Typically, strains should be grown 1 to 2 days
in raffinose, followed by growth for 2 days in
galactose or glucose. An additional 1 to 2 days
may be necessary to allow growth of “pop-out”
strains on 5-FOA plates.
Vectorette PCR and subsequent DNA sequence analysis may be performed within a
similar time frame. In preparation for PCR, a
total of 2 to 3 days may be required to synthesize
appropriate primers and isolate genomic DNA.
Anchor bubbles may be ligated to genomic
DNA in 1 day. PCR itself requires only a few
hours. PCR products may be analyzed and
recovered in ∼6 hr. DNA sequencing reactions
can be completed in 1 day, and results may be
viewed the next morning. In total, an initial
attempt at vectorette PCR may require ∼5 days;
however, subsequent vectorette PCR may be
accomplished in only 3 days. Potential stopping
points for both procedures have been indicated
at appropriate steps in the respective protocols.
Literature Cited
Burns, N., Grimwade, B., Ross-Macdonald, P.B.,
Choi, E.-Y., Finberg, K., Roeder, G.S., and Snyder, M. 1994. Large-scale analysis of gene expression, protein localization and gene disruption in Saccharomyces cerevisiae. Genes Dev.
8:1087-1105.
Chen, D.C., Yang, B.C., and Kuo, T.T. 1992. Onestep transformation of yeast in stationary phase.
Curr. Genet. 21:83-84.
Devine, S.E. and Boeke, J.D. 1994. Efficient integration of artificial transposons into plasmid targets in vitro: a useful tool for DNA mapping,
sequencing and genetic analysis. Nucleic Acids
Res. 22:3765-3772.
Genome-Wide
Transposon
Mutagenesis in
Yeast
Garfinkel, D.J. and Strathern, J.N. 1991. Ty mutagenesis in Saccharomyces cerevisiae. Methods
Enzymol. 194:342-361.
Hoekstra, M.F., Seifert, H.S., Nickoloff, J., and Heffron, F. 1991. Shuttle mutagenesis: bacterial
transposons for genetic manipulation in yeast.
Methods Enzymol. 194:329-342.
Ji., H., Moore, D.P., Blomberg, M.A., Braiterman,
L.T., Voytas, D.F., Natsoulis, G., and Boeke, J.D.
1993. Hotspots for unselected Ty1 transposition
events on yeast chromosome III are near tRNA
genes and LTR sequences. Cell 73:1007-1018.
Riley, J., Butler, R., Ogilvie, D., Finniear, R., Jenner,
D., Powell, S., Anand, R., Smith, J.C., and Markham, A.F. 1990. A novel, rapid method for the
isolation of terminal sequences from yeast artificial chromosome (YAC) clones. Nucleic Acids
Res. 18:2887-2890.
Ross-Macdonald, P., Sheehan, A., Roeder, G.S., and
Snyder, M. 1997. A multipurpose transposon
system for analyzing protein production, localization, and function in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. U.S.A. 94:190-195.
Ross-Macdonald, P., Coelho, P., Roemer, T., Agarwal, S., Kumar, A., Cheung, K.-H., Jansen, R.,
Sheehan, A., Symoniatis, D., Umansky, L., Nelson, K., Iwasaki, H., Hager, K., Gerstein, M.,
Miller, P., Roeder, G.S., and Snyder, M. 1999.
Large-scale analysis of the yeast genome by
transposon tagging and gene disruption. Nature
402:413-418.
Seifert, H.S., Chen, E.Y., So, M., and Heffron, F.
1986. Shuttle mutagenesis: A method of
transposon mutagenesis for Saccharomyces
cerevisiae. Proc. Natl. Acad. Sci. U.S.A. 83:735739.
Key References
Kumar, A., Cheung, K.-H., Ross-MacDonald, P.,
Coelho, P.S.R., Miller, P., and Snyder, M. 2000.
TRIPLES: A database of gene function in S.
cerevisiae. Nucleic Acids Res. 28:81-84.
Provides a helpful explanation of resources freely
available from the authors’ web site (see Internet
Resources).
Seifert et al., 1986. See above.
An early application of Tn3-based shuttle mutagenesis to Saccharomyces cerevisiae.
Ross-Macdonald et al., 1997. See above.
Provides an in-depth description of multifunctional
Tn3-minitransposons used in this unit.
Ross-Macdonald et al., 1999. See above.
Presents an extensive application of Tn3-mediated
shuttle-mutagenesis towards functional genomics in
yeast, with protocols for the genome-wide analysis
of disruption phenotypes, gene expression, and protein localization.
13.3.14
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Current Protocols in Molecular Biology
Internet Resources
http://ygac.med.yale.edu
Many strains and reagents used in this protocol
(including all transposon-insertion libraries) may
be requested from this, the authors’ web site.
Contributed by Anuj Kumar and
Michael Snyder
Yale University
New Haven, Connecticut
Saccharomyces
cerevisiae
13.3.15
Current Protocols in Molecular Biology
Supplement 51
EMS and UV Mutagenesis in Yeast
UNIT 13.3B
1
Fred Winston
1
Harvard Medical School, Boston, Massachusetts
ABSTRACT
Many fundamental biological processes have been elucidated by the isolation and analysis
of mutants that are defective in such processes. Therefore, the methods to generate
mutants are of great importance in model organisms. This unit describes two protocols
for mutagenesis of yeast—using ethyl methanesulfate (EMS) and ultraviolet (UV) light.
Each of these methods has been used successfully for many years. Curr. Protoc. Mol.
C 2008 by John Wiley & Sons, Inc.
Biol. 82:13.3B.1-13.3B.5. Keywords: yeast r mutagenesis r EMS r UV
INTRODUCTION
Because spontaneous mutations occur at a low rate, yeast cells are often treated with
mutagens to increase the frequency of mutants. Two common mutagens of yeast cells are
ethyl methanesulfonate (EMS) and ultraviolet (UV) light. Mutagenesis can increase the
frequency of mutation up to 100-fold per gene without excessive killing of the cells and
without a significant frequency of double mutants. EMS and UV may produce different
spectra of mutants, but generally only one type of mutagenesis is necessary to generate
a sufficient number of mutants to study.
This unit presents protocols for both EMS and UV mutagenesis of yeast cells. Cultures of
the desired yeast strain are treated with the mutagen, and the effectiveness of the mutagen
is measured by determining the frequency of an event for which there exists a genetic
selection (e.g., canavanine resistance). Mutagenized cells can then be screened for any
phenotype of interest, including auxotrophies, cold sensitivity, and radiation sensitivity.
As an example, the Basic Protocol includes steps for screening for mutants that are
temperature-sensitive for growth. Once a gene of interest has been identified by such
procedures, it can be cloned, mutagenized, and manipulated in other ways to study its
function in greater detail (UNITS 13.8 & 13.10).
NOTE: All solutions, plasticware, glassware, and velveteens coming into contact with
yeast cells must be sterile.
MUTAGENESIS USING ETHYL METHANESULFONATE (EMS)
Materials
BASIC
PROTOCOL
Desired yeast strain
YPD medium and plates (UNIT 13.1)
Sterile water
0.1 M sodium phosphate buffer, pH 7.0 (see recipe)
Ethyl methanesulfonate (EMS; Kodak)
5% (w/v) sodium thiosulfate (Sigma), autoclaved for sterility
13 × 100–mm culture tube
Vortex
30◦ C incubator with rotating platform
Canavanine plates (UNIT 13.1)
Yeast
Current Protocols in Molecular Biology 13.3B.1-13.3B.5, April 2008
Published online April 2008 in Wiley Interscience (www.interscience.wiley.com).
DOI: 10.1002/0471142727.mb1303bs82
C 2008 John Wiley & Sons, Inc.
Copyright 13.3B.1
Supplement 82
Additional reagents and equipment for growing cells, determining cell density, and
replica plating (UNIT 13.2) and for storage of strains (UNIT 13.1)
CAUTION: EMS is a dangerous mutagen. All solutions, plasticware, and glassware that
come into contact with EMS should be rinsed with 5% sodium thiosulfate to inactivate
the EMS.
Grow and mutagenize cells
1. Grow an overnight culture of the desired yeast strain (UNIT 13.1) in 5 ml YPD medium
at 30◦ C.
2. Determine the density of cells (UNIT 13.2) in the culture and record this number.
Adjust concentration to ∼2 × 108 cells/ml if necessary. Transfer 1 ml of the culture
to a sterile microcentrifuge tube.
3. Pellet cells in a microcentrifuge ∼5 to 10 sec at maximum speed, room temperature.
Discard supernatant and resuspend in 1 ml sterile water. Repeat wash. After the
second wash, resuspend cells in 1.5 ml sterile 0.1 M sodium phosphate buffer,
pH 7.0.
4. Add 0.7 ml cell suspension to 1 ml buffer in a 13 × 100–mm culture tube. Save
remaining cells on ice for a control.
5. Add 50 µl EMS to the cells and disperse by vortexing. Place on a rotating platform
and incubate 1 hr at 30◦ C.
EMS treatment should cause ∼40% of the cells to be killed.
6. Transfer 0.2 ml of the treated cell suspension to a culture tube containing 8
ml sterile 5% sodium thiosulfate, which will stop the mutagenesis by inactivation of EMS. If cells are to be stored before plating, pellet in a tabletop centrifuge 5 min at 3000×g, 4◦ C), resuspend in an equal volume of sterile water,
and store at 4◦ C.
Determine effectiveness of mutagenesis
7. Plate 0.1 ml mutagenized cells directly on each of two canavanine plates. As
controls, plate 0.1 ml nonmutagenized cells (from step 4) on duplicate canavanine
plates. Incubate at 30◦ C until colonies form (∼2 to 4 days).
8. Calculate the relative levels of canavanine-resistant mutants in the mutagenized and
nonmutagenized cultures.
The percentage survival can be calculated by comparing the number of cells in the initial
culture (step 2) and the number of viable cells following EMS treatment (calculated from
the number of colonies obtained in step 7).
Identify temperature-sensitive mutants
9. Dilute the EMS-treated cells from step 6 with sterile water to obtain 100 to 200
viable cells per plate.
A dilution factor of 1:1000 may be sufficient, but this will depend on the initial concentration of cells used.
10. Plate 0.1 and 0.2 ml of the diluted cells on separate sets of YPD plates, using ten
plates in each set. Incubate all plates 3 to 4 days at room temperature (23◦ C).
EMS and UV
Mutagenesis in
Yeast
For identification of other types of mutants, plates can be incubated at other temperatures. For example, if screening for auxotrophs, incubate plates at 30◦ C, the optimal
temperature for growth of yeast cells. In addition, the number of plates used may vary,
depending upon the frequency of the desired mutant class. Up to 20,000 colonies (l00
plates, each with 200 colonies) can be screened if necessary.
13.3B.2
Supplement 82
Current Protocols in Molecular Biology
11. Choose at least ten YPD plates that contain 100 to 200 colonies per plate. Replicaplate (UNIT 13.2) each to two fresh YPD plates. For temperature-sensitive mutants,
incubate one YPD plate per set at 37◦ C and one at room temperature (23◦ C),
overnight.
The 23◦ C plate serves as a positive control for growth, as some temperature-sensitive
mutants may grow poorly at 30◦ C.
Be certain that each plate is numbered and has an orientation symbol on the back. In
general, one of the two replica plates should be permissive for mutant growth (typically
a YPD plate). The other plate should represent conditions that do not permit growth of
the desired class of mutants.
12. Compare the 37◦ C plate with the 23◦ C and 30◦ C plates. Any colonies that failed
to grow at 37◦ C are candidates for temperature-sensitive mutants. To recheck, pick
corresponding colonies from YPD plates incubated at 23◦ C and streak for single
colonies on fresh YPD plates. On each plate, also streak the parental control strain
for single colonies. Incubate the plates at 23◦ C until colonies form (2 to 4 days).
Six to eight strains can be purified on each YPD plate.
13. After single colonies form, replica-plate (UNIT 13.2) the YPD plates with the
temperature-sensitive candidates to two YPD plates as before. Incubate one plate
at 37◦ C and the other at room temperature for 1 day.
14. Record the growth response. For each candidate that is reproducibly temperaturesensitive, place into permanent storage (UNIT 13.1).
MUTAGENESIS USING UV IRRADIATION
Additional Materials (also see Basic Protocol)
ALTERNATE
PROTOCOL
UV germicidal light bulb (Sylvania G15T8; 254 nm wavelength) or Stratagene UV
Crosslinker
UV dosimeter (optional)
23◦ C incubator
CAUTION: Wear safety glasses to protect eyes from UV light.
1. Grow an overnight culture of the desired yeast strain (UNIT 13.2) in 5 ml YPD medium
at 30◦ C.
2. Pellet 1 ml of cells in a microcentrifuge ∼5 to 10 sec at maximum speed, room
temperature and discard supernatant. Resuspend in 1 ml sterile water and repeat
wash. After the second wash, resuspend in 1 ml sterile water.
3. Determine the cell density and record this number. Adjust to 2 × 108 cells/ml if
necessary.
4. Make serial dilutions of the culture in sterile water so that each plate has 200 to
300 viable cells. Plate 0.1 and 0.2 ml on YPD plates as described in step 10 of the
Basic Protocol.
5. Irradiate all but two plates from each set with UV light using a dosage of
300 ergs/mm2 . There should be ∼40% to 70% survival. The nonirradiated plates
will serve as controls to determine the degree of killing by the UV light.
Light from the UV germicidal bulb can be measured using a UV dosimeter. From this
measurement, the proper length of time can be calculated for irradiation to attain 300
ergs/mm2 . Alternatively, the proper time of irradiation can be empirically determined
by measuring the time that results in 40% to 70% survival.
Yeast
13.3B.3
Current Protocols in Molecular Biology
Supplement 82
Most UV lights should be warmed up for ∼20 min before use. In addition, petri plate lids
block transmission of UV light; therefore, be sure to remove them prior to irradiation.
Alternatively, a Stratagene UV Crosslinker, commonly used for cross-linking DNA or
RNA to membranes, can be used. As for a germicidal lamp, the proper time should be
determined empirically.
6. As a control to determine the degree of UV mutagenesis, plate 0.1 ml of the
original culture on duplicate canavanine plates and irradiate one plate for the time
determined in step 5. From the number of canavanine-resistant colonies, calculate
the frequency of mutations with and without UV irradiation.
As described in the Basic Protocol, the degree of mutagenesis can be determined by
measuring traits other than canavanine resistance.
7. Incubate plates from step 4 at 23◦ C until colonies form (∼2 to 4 days).
8. Screen colonies for temperature-sensitive mutants as described in steps 9 to 14 of
the Basic Protocol.
REAGENTS AND SOLUTIONS
Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see
APPENDIX 2; for suppliers, see APPENDIX 4.
Sodium phosphate buffer, 0.1 M, pH 7.0
Prepare 1 M solutions of Na2 HPO4 and NaH2 PO4 . Mix 5.77 ml Na2 HPO4 with
4.23 ml NaH2 PO4 and add water to 100 ml (the pH will be 7.0). Autoclave for
sterility. Store up to 1 year at room temperature.
COMMENTARY
Background Information
EMS and UV
Mutagenesis in
Yeast
13.3B.4
Supplement 82
Mutagenesis of yeast cells is an important
genetic technique to increase the frequency
and spectrum of mutants obtained. Even when
mutants arise spontaneously through genetic
selection, treatment with a chemical mutagen
or irradiation can alter the types of mutants
obtained. EMS is known to cause primarily
GC to AT transversions. UV light has been reported to cause a wide spectrum of mutational
changes, including transitions and transversions (Prakash and Sherman, 1973; Coulondre
and Miller, 1977).
Mutagens will generally alter only one
DNA strand at any mutagenized site. This
should create a double-stranded molecule that
is heterozygous, possibly resulting in a yeast
colony of mixed genotype. In practice, this
is rarely observed. Evidence exists that mismatch repair may play a role in the degree and
spectrum of mutagenesis (Muster-Nassal and
Kolodner, 1986; Siede and Eckardt-Schupp,
1986).
The degree of mutagenesis can be monitored by measuring any easily detectable mutational event. Instead of using canavanine resistance as in the Basic Protocol, one could,
for example, measure α-aminoadipate utilization (UNIT 13.1) or the reversion of a revertable
auxotrophic mutation present in the parental
strain.
Critical Parameters
The degree of mutagenesis is a critical factor. Too little mutagenesis will not sufficiently
increase the frequency of mutants in a population of cells; too much mutagenesis will result
in multiple mutations. Since some mutagens,
such as nitrosoguanidine, create multiple mutations in a nonrandom fashion (Botstein and
Jones, 1969), double mutants may not be easily discovered by standard genetic analysis.
Following isolation, any mutants of interest should be subjected to basic genetic analysis (Rose et al., 1990). The first step in this
analysis should be a genetic cross (UNIT 13.2)
to demonstrate that the mutant phenotype is
caused by a mutation in a single gene (i.e.,
that the mutant phenotype segregates 2:2 in
tetrads). If a large number of mutants are isolated, the number of genes identified can be
determined by complementation and linkage
analysis (Rose et al., 1990).
Anticipated Results
Under optimal conditions of mutagenesis
as described in this unit, the frequency of
canavanine-resistant mutants should be at least
Current Protocols in Molecular Biology
l00-fold greater after mutagenesis. This result
is important in order to know that the mutagenesis has worked properly, since the frequency of obtaining the desired class of mutant is likely to be unknown. The frequency
of finding a particular class of mutant will depend entirely upon the nature of the mutation
sought. For example, random temperaturesensitive mutants will be found at a higher frequency than temperature-sensitive mutations
that cause cell-cycle arrest in G2.
Time Considerations
Mutagenizing cells with EMS takes ∼2 hr.
Plating the cells requires ∼30 min, with colony
formation in 3 to 4 days. Mutagenizing cells
by UV irradiation requires ∼1 hr to prepare
serial dilutions and plate cells. Once the plates
have been irradiated, colony formation takes 2
to 4 days. For both methods, identification of
temperature-sensitive mutants will require an
additional 4 to 8 days.
Literature Cited
Botstein, D. and Jones, E.W. 1969. Nonrandom
mutagenesis of the Escherichia coli genome
by nitrosoguanidine. J. Bacteriol. 98:847848.
Coulondre, C. and Miller, J.H. 1977. Genetic studies of the lac repressor. IV. Mutagenic specificity
in the lacI gene of Escherichia coli. J. Mol. Biol.
117:577-606.
Muster-Nassal, C. and Kolodner, R. 1986. Mismatch correction catalyzed by cell-free extracts
of Saccharomyces cerevisiae. Proc. Natl. Acad.
Sci. U.S.A. 83:7618-7622.
Prakash, L. and Sherman, F. 1973. Mutagenic specificity: Reversion of iso-1-cytochrome c mutants
of yeast. J. Mol. Biol. 79:65-82.
Rose, M.D., Winston, F., and Hieter, P. 1990.
Laboratory Course Manual for Methods in Yeast
Genetics. Cold Spring Harbor Laboratory, Cold
Spring Harbor, New York.
Siede, W. and Eckardt-Schupp, F. 1986. A mismatch repair-based model can explain some features of UV mutagenesis in yeast. Mutagenesis
1:471-474.
Yeast
13.3B.5
Current Protocols in Molecular Biology
Supplement 82
SECTION II
UNIT 13.4
YEAST VECTORS
Yeast Cloning Vectors and Genes
resistance. In contrast, the cloned yeast gene
present on the plasmid is a copy of a gene that
is present in the yeast genome as well. Thus,
this gene functions as a dominant selectable
marker only when the recipient yeast cell has a
recessive mutation in the corresponding chromosomal copy of the cloned gene. Many of
these cloned yeast genes encode functions involved in biosynthetic pathways of yeast and
are capable of complementing certain mutations in similar biosynthetic pathways of E. coli
(e.g., the cloned URA3 gene of yeast can complement ura3− mutations of yeast as well as
mutations in the pyrF gene of E. coli). The
availability of bacterial mutations that can be
complemented by these yeast genes can greatly
simplify plasmid constructions, allowing genetic screening for the expected recombinant
plasmid.
Table 13.4.2 presents characteristics of a
number of cloned yeast genes, along with
This unit describes some of the most commonly used yeast vectors, as well as the cloned
yeast genes that form the basis for these plasmids. Yeast vectors can be grouped into five
general classes, based on their mode of replication in yeast: YIp, YRp, YCp, YEp, and YLp
plasmids. With the exception of the YLp plasmids (yeast linear plasmids), all of these plasmids can be maintained in E. coli as well as in
S. cerevisiae and thus are referred to as shuttle
vectors.
Table 13.4.1 summarizes the general features of a number of these vectors, including
the phenotypes that allow selection in either
E. coli or yeast (or both). These plasmids contain two types of selectable genes, both of
which can confer a dominant phenotype: plasmid-encoded drug-resistance genes (UNIT 1.5)
and cloned yeast genes. The drug resistance
marker is dominant because the recipient (bacterial) cell does not encode a gene for drug
Table 13.4.1
Yeast Plasmids and their Selectable Markers
Plasmid
Size
Yeast
E. coli
replicon replicon
YIp5
5541 bp
pMB1
none
YRp7
5816 bp
pMB1
ARS1
YRp17
7002 bp
pMB1
ARS1
YEp13
10.7 kb
pMB1
2µm
YEp24
7769 bp
pMB1
25µm
YCp19
10.1 kb
pMB1
ARS1
YCp50
7.95 kb
pMB1
ARS1
YLp21
55 kb
none
ARS1
pYAC3
11.4 kb
pMB1
ARS1
2µm
6318 bp
none
2µm
Phenotypes
selectable
in E. coli
Phenotypes
selectable
in yeast
Reference
Ampr, Tetr,
PyrF+
Ampr, Tetr,
TrpC+
Ampr, Tetr,
PyrF+, TrpC+
Ampr, Tetr,
LeuB+
Ampr;
Tetr in some
constructions;
PyrF+
Ampr, PyrF+,
TrpC+
Ampr, Tetr,
PyrF+
n.a.
Ura+
Struhl et al., 1979
Trp+
Struhl et al., 1979
Ura+, Trp+
Stinchcomb et al.,
1982
Broach et al.,
1979
Botstein et al.,
1979
Ampr, Tetr,
TrpC+, PyrF+,
HisB+
n.a.
Trp+, Ura+,
His+
Yeast Cloning
Vectors and Genes
13.4.1
Supplement 21
Contributed by Victoria Lundblad
Current Protocols in Molecular Biology (1992) 13.4.1-13.4.10
Copyright © 2000 by John Wiley & Sons, Inc.
Leu+
Ura+
Ura+, Trp+
Ura+
Trp+, His+
none
Stinchcomb et al.,
1982
Rose et al., 1987
Murray and
Szostak, 1983
Burke et al., 1987
Hartley and
Donelson, 1980
Table 13.4.2
Cloned Yeast Genes
Length of
sequenced
fragment (bp)
Map
position
in yeast
genome
ARG4
ARS1
CAN1d
2296
1453c
n.s.
CEN3
CEN4
CYH2e
GAL1GAL10
regulatory
region
HIS3
Selectable
phenotype
of wild type
in yeast
Selectable
phenotype
in E. coli
E. coli
straina
8R
4R
5L
ArgH+
none
none
JA209
n.a.
n.a.
Arg+
none
Cans
627
2095
1393
907
3
4
7L
2R
none
none
none
none
n.a.
n.a.
n.a.
n.a.
none
none
viabilityf
none
1822
15R
HisB+
BA1
His+
LEU2
LYS2
TCM1g
TRP1
2230
n.s.
1529
1453c
3L
2R
15R
4R
LeuB+
none
none
TrpC+
JA300
n.a.
n.a.
JA300
Leu+
Lys+
viabilityf
Trp+
URA3
1170
5L
PyrF+
DB6656
Ura+
Yeast
gene
Common
nonreverting
mutant
alleles
Selectable
phenotype of
mutant alleles
in yeastb
none
n.a.
can1-100
can1-11
n.a.
n.a.
none
none
canavanine
sulfater
none
none
cycloheximider
n.a.
YSCCEN3
YSCCEN4
YSCRPL29
YSCGAL
none
SCHIS3Y
none
α-aminoadipater
tricoderminr
none
YSCLEU2
n.a.
YSCRP13
YSCTRP1
5-fluoroorotic
acidr
YSCODCD
n.a.
his3-∆1
his3-200
leu2-3,112
lys2-∆1
n.a.
trp1-289
trp1-∆901
ura3-52
ura3-∆1
GenBank
file name
YSCARG4
YSCTRP1
YSCCAN1
n.s. = not sequenced; n.a. = not applicable.
aThe genotypes and references for these strains can be found in the legends to Figures 13.4.1, 13.4.2, and 13.4.6, except for JA209 (argH1 metE xyl
trpA36 recA56 strr; Clarke and Carbon, 1978).
bSelection for α-aminoadipate resistance produces both lys2− and lys5− mutants. Selection for 5-FOA resistance generates both ura3− and ura5− mutants.
cThis 1453-bp fragment contains both the TRP1 and the ARS1 genes.
dThe wild-type CAN1 gene encodes dominant sensitivity to the arginine analog canavanine sulfate.
eThe sensitive (wild-type) and resistant alleles of the CYH2 gene are codominant.
fThe CYH2 and TCM1 genes encode ribosomal proteins, which are required for viability, but which can be mutated to confer resistance to cycloheximide
or tricodermin, respectively.
gThe resistant allele of TCM1 is dominant to the wild-type sensitive allele.
mutants of bacteria and yeast in which the
wild-type cloned gene can be selected. For two
yeast genes, a positive selection for mutant
alleles also exists: ura3− cells can be selected
on plates containing the drug 5-fluoroorotic
acid (5-FOA), whereas lys2− cells can be selected on α-aminoadipate plates (see UNIT 13.2
for recipes for these two types of plates).
The nomenclature of different classes of
yeast vectors, as well as details about their mode
of replication in yeast are described below.
PLASMID NOMENCLATURE
YIp plasmids (yeast integrating plasmids)
contain selectable yeast genes but lack sequences that allow autonomous replication of
the plasmid in yeast. Instead, transformation of
yeast occurs by integration of the YIp plasmid
into the yeast genome by recombination between yeast sequences carried on the plasmid
and homologous sequences in the yeast
genome. This recombination event results in a
tandem duplication of the yeast sequences that
bracket the rest of the plasmid DNA. If the YIp
plasmid contains an incomplete portion of a
cloned gene, this technique can be used to
create a gene disruption (see UNIT 13.10). The
reversal of the integration process occurs at a
low frequency (about 0.1% to 1% per generation), with excision of the integrated plasmid
occurring by recombination between duplicated yeast sequences. The frequency of transformation of YIp plamids is only 1 to 10 transformants/µg DNA, but transformation frequency can be increased 10- to 1000-fold by
linearizing the plasmid within yeast sequences
Saccharomyces
cerevisiae
13.4.2
Current Protocols in Molecular Biology
Supplement 26
Yeast Cloning
Vectors and Genes
that are homologous to the intended site of
integration on the yeast chromosome. Linearization also directs the integration event to the
site of the cleavage, which is useful when several different homologous yeast sequences are
present on the plasmid.
Three classes of yeast vectors are circular
plasmids capable of extrachromosomal replication in yeast. YRp plasmids (yeast replicating
plasmids) contain sequences from the yeast
genome which confer the ability to replicate
autonomously. These autonomous replication
sequences (ARS) have been shown to be chromosomal origins of replication. YRp plasmids
have high frequencies of transformation (103 to
104 transformants/µg DNA), but transformants
are very unstable both mitotically and meiotically. Despite the fact that ARS-containing plasmids replicate only once during the cell cycle,
YRp plasmids can be present in high copy
number (up to 100 copies per plasmid-bearing
cell, although the average copy number per cell
is 1 to 10). During mitosis, both instability and
high copy number are due to a strong bias to
segregate to the mother cell, to the extent that
as few as 5% to 10% of cells grown selectively
still retain the plasmid.
Incorporation of DNA segments from yeast
centromeres (CEN elements) into YRp plasmids, to generate vectors called YCp plasmids
(yeast centromeric plasmids), greatly increases
plasmid stability during mitosis and meiosis.
Such plasmids—present in 1 to 2 copies per
cell—have a loss rate of approximately 1% per
generation and show virtually no segregation
bias. During meiosis, CEN plasmids behave
like natural chromosomes, generally segregating in a 2+:2− ratio.
The last class of circular replicating plasmids, YEp vectors (yeast episomal plasmids),
contain sequences from a naturally occurring
yeast plasmid called the 2µm circle. These 2µm
sequences allow extrachromosomal replication
and confer high transformation frequencies
(∼104 to 105 transformants/µg DNA). These
plasmids are commonly used for high-level
gene expression in yeast, due to their ability to
be propagated relatively stably through mitosis
and meiosis in high copy number. YEp vectors
vary in the portion of 2µm DNA that they carry,
although most carry only the sequences essential for autonomous replication. If the 2µm-encoded REP1 and REP2 functions are present
(either on the YEp plasmid or due to the presence of endogenous 2µm circles), transformants are relatively stable and present in high
copy number (20 to 50 copies). In the absence
of REP functions, transformants are much more
unstable, with a segregation bias and copy number similar to those observed with YRp plasmids. In the 2µm circle, a highly efficient sitespecific recombination event occurs between
two perfect 599-bp inverted repeats, mediated
by the 2µm–encoded FLP gene. Most YEp
plasmids carry at least one copy of this repeat;
thus, FLP-mediated recombination between
YEp vectors and other plasmids carrying one
or more of these repeats (either the endogenous 2µm plasmid or other shuttle vectors)
can result in a variety of recombinant plasmid
multimers.
YLp plasmids (yeast linear plasmids) contain certain G-rich repeated sequences at their
termini which function as telomeres and allow
the plasmid to replicate as a linear molecule. In
yeast, the telomeric sequence consists of tandem repeats of the sequence 5′(dG1-3dT)3′.
Very short CEN-containing YLp plasmids (10
to 15 kb) are unstable and present in high copy,
due to random segregation during mitosis. Increasing the size to 50 to 100 kb produces YLp
vectors that disjoin from each other in a manner
similar to that of natural chromosomes, resulting in a copy number of about one per cell.
However, these artificial chromosomes—
which are lost at a rate of 10−2 to 10−3 per cell
division—are still ∼100-fold less stable than a
natural yeast chromosome.
MAPS OF SELECTED PLASMIDS
AND GENES
Restriction maps and a brief description of
selected plasmid vectors from several of the five
general classes are presented in Figures 13.4.1,
13.4.2, 13.4.3, 13.4.4, 13.4.5, 13.4.6, and
13.4.7. These plasmids were chosen because
they are used by many investigators and are
generally applicable for a wide variety of purposes. However, where different selectable
markers or unique restriction sites may be required, the reader is referred to two reviews on
vector systems used in yeast (Pouwels et al.,
1985; Parent et al., 1985). In addition, a method
for constructing new plasmids in vivo in yeast
has been described and employed to construct
an extended series of new YRp, YCp, and YEp
plasmids (Ma et al., 1987). Yeast shuttle vectors have also been constructed that are derived from either pUC18 or the Bluescript
plasmids, providing a greater variety of unique
cloning sites and allowing both identification
of recombinant plasmids by screening for alpha-
13.4.3
Supplement 26
Current Protocols in Molecular Biology
EcoRI
1/5541 Clal27
HindIII 32
AatII 5471
Nhel 232
Sspl 5353
BamHI 378 SphI 569
EcoNI 629
Pvul 4919
Sall 654
XmaIII 942
r
NruI 977
Apr
Tet
bacterial DNA
yeast DNA
YIp5
5541 bp
pMB1
ori
URA3
1030
227
380 Gsul
415 EcoRV
432 Ncol
494 Ksp 632I
492 Asu ll
501 Bbv ll
539 Scal
553 Xcal
589 Bsml
593 Dral
595 Dral
605 Apal
663 Stul
729 Gsul
737 Ksp632l
790 Ksp632l
833 Dralll
901 Eco31l
908 Alw Nl
922 Ksp632l
URA3
140 Ndel
209 Pstl
212 BspMl
Hindlll
URA3
Hindlll
Nsil 2488
Xmal 2540
BspMII 2845
Ball 2627 Smal 2542
1054 Nsi l
1058 Xcal
1106 Xmal
1108 Smal
Tth111l 3403
Pvull 3249
Ncol 1866
AsuII 1926
Apal 2039
Stul 2097
Figure 13.4.1 YIp5. YIp5 contains the 1.1-kb HindIII URA3 gene cloned into the AvaI site of
pBR322 via the addition of poly(dG-dC) tails (Struhl et al., 1979). Since this plasmid does not contain
a yeast origin of replication, transformants occur by integration into the yeast genome at the URA3
locus; the frequency of transformation can be increased by linearization of the plasmid within
the URA3 insert. The complete nucleotide sequence is available from the Vecbase database (file
name: Vecbase.Yip5) and a detailed restriction map can be found in the New England Biolabs
catalog.
The URA3 gene encodes orotidine-5′-phosphate (OMP) decarboxylase, a 267-amino-acid
protein required for uracil biosynthesis. The map shown is the gene from strain +D4, which can be
expressed in E. coli without an external bacterial promoter (Rose et al., 1984). Loss of URA3+
function can be directly selected using 5-FOA: ura3− cells are resistant to 5-FOA, whereas 5-FOA
is toxic to cells synthesizing the URA3 gene product (Boeke et al., 1984). This negative selection
has been exploited in a variety of gene replacement schemes, discussed in UNIT 13.10. The URA3
gene can also complement mutations in the pyrF gene in E. coli using strain DB6656 (pyrF::Mu,
lacZam trpam hsrk− hsmk+; Bach et al., 1979).
complementation of the lacZÄM15 mutation of
E. coli (see UNIT 1.4) and the ability to produce
single-stranded DNA (Hill et al., 1986; Elledge
and Davis, 1988).
Finally, a vector system has been designed
using yeast artificial chromosome (pYAC)
plasmids, allowing direct cloning into yeast of
contiguous stretches of DNA up to 400 kb
(Burke et al., 1987). The circular pYAC plasmids (without inserts) can replicate in E. coli.
In vitro digestion of the pYAC vector, ligation
to exogenous DNA, and direct transformation
of the subsequent linear molecules (with
telomeric sequences at each termini) into yeast
generate a library that can then be screened by
standard techniques.
Saccharomyces
cerevisiae
13.4.4
Current Protocols in Molecular Biology
Supplement 7
bacterial DNA
yeast DNA
AatII 5743
SspI 5625
1/5816
Dralll 26
Bgl ll 601
Stul 624
ScaI 5301
PvuI 5191
ARS1
Bst XI 1053
AocI 1109
Apr
Eco31I 4882
XbaI 1267
TRP1
PmaCI 1391
CIaI 1477
YRp7
5816 bp
AIw NI 4344
pMB1
ori
Tet r
615
TRP1 ARS1
1453
BamHI 1828
SphI 2019
EcoNI 2079
SaII 2104
NdeI 3751
XmaIII 2392
Tth111I 3675
NruI
2427
PvuII 3521
BsmI 2812
StyI 2822
BspMII 3117
BaII 2899 AvaI 2878
ARS1
EcoRI
1389 ApaLI
1417 DraIII
987 Ks p6321
1037 NheI
1069 NaeI
813 ApaLI
827 Ps tI
829 StuI
852 BgI II
615 Hin dIII
535 BspMI
387 Eco RV
400 Bs tXI
344 AccI
186 XbaI
777
103
62 Pma CI
69 XcaI
Ec oRI
TRP1
Figure 13.4.2 YRp7. This plasmid contains the 1453-bp EcoRI TRP1 ARS1 fragment from S.
cerevisiae inserted into the EcoRI site of pBR322 (Struhl et al., 1979).
The TRP1 RI circle is a derivative of YRp7 containing only the 1453-bp TRP1 ARS1 EcoRI
fragment. This plasmid is mitotically and meiotically unstable, but is present in 100 to 200 copies
per plasmid-bearing cell in both cir+ and ciro strains.
A genomic plasmid bank has been constructed by inserting size-selected Sau3A partial fragments into the BamHI site of YRp7 (Nasmyth and Reed, 1980).
The TRP1 ARS1 1453-bp EcoRI fragment contains both the TRP1 gene, encoding N-(5′-phosphoribosyl)-anthranilate isomerase, and the autonomous replication sequence ARS1. The intact
TRP1 gene can complement mutations in the trpC gene of E. coli, using E. coli JA300 (thr1 leuB6
thi1 thyA trpC1117 hsrk− hsmk− strr; Tschumper and Carbon, 1982). The chromosomal replicator
ARS1 lies between positions 615 and 1453 (on a HindIII-EcoRI fragment) and is composed of three
domains. Domain A contains an 11-bp core sequence (position 857 to 867) consisting of a
consensus sequence found in many other ARS elements and which is essential for ARS1 function.
Domains B and C flank Domain A and are relatively AT-rich regions that contribute to, but are not
essential, for ARS function.
Yeast Cloning
Vectors and Genes
13.4.5
Supplement 7
Current Protocols in Molecular Biology
1/7769
BcII 304
Aat II 7699
XbaI 703
bacterial DNA
yeast DNA
SnaBI 1044
ampr
2µm origin
pMB1
ori
HpaI 1686
YEp24
7769 bp
EspI 2125
Tth111I 5631
PvuII 5477
ClaI 2267
URA3
NcoI 2704
ApaI 2877
tetr (?)
BspMII 5073
XmaI 3378
XmaIII 4348
SmaI 3380
SaII 4060
NheI 3638
Eco NI 4035
BamHI 3784
SphI 3975
Figure 13.4.3 YEp24. YEp24 has the 2.2-kb EcoRI fragment of the B form of the 2 µm plasmid
and the 1.1-kb HindIII URA3 gene inserted into the EcoRI and HindIII sites, respectively, of pBR322
(Botstein et al., 1979). The expression of the tetr gene is variable among different isolates of this
plasmid. YEp24 is mitotically stable in cir+ strains at a copy number of about 20 but is unstable in
ciro strains. The complete sequence of YEp24 is available from the Vecbase database (file name:
Vecbase.Yep24) and a detailed restriction map can be found in the New England Biolabs catalog.
SwaI 6175
Bsu 36I 5603
Hgi EII 5599
SphI 5568
EagI 5424
BcgI 244
MscI 264
ClaI (6284)
BcII 304
NdeI 339
FLP
IR2
BspMI 5319
BsiI 1324
HgiAI 1422
NcoI 5024
StyI 5024
REP1
2 µm
6318 bp
AseI 1664
REP2
Alw NI 1888
Esp 3I 1913
IR1
BanII 4387
ApaI 4387
D protein
Bsr BI 2463
NruI 2558
TaqII 2581
SnaBI 3606
HpaI 2964
PstI 2652
Figure 13.4.4 2ìm plasmid. The 2µm circle is a naturally occurring DNA plasmid found in almost
all strains of S. cerevisiae, with a copy number of ∼20 to 80. The plasmid exists in two different forms,
A and B (the former is shown above), due to intra-molecular recombination between two perfect
599-bp inverted repeats. Strains that carry this plasmid are called cir+; strains missing the plasmid
cir0 have been identified or isolated (see UNIT 13.9). It is extremely stable mitotically, with a
spontaneous loss rate in haploid cells of 10−4 per generation; during meiosis the plasmid is
transmitted to all four spore products. The plasmid has been completely sequenced (Hartley and
Donelson, 1980) and the sequence is available from GenBank (Plant: yscplasm).
Saccharomyces
cerevisiae
13.4.6
Current Protocols in Molecular Biology
Supplement 30
EcoRI
1/ 7950
Hin dIII 32
Aat II 7880
BamHI 378
SphI 569
PvuI 7325
SaII 654
XmaIII 942
NruI 977
bacterial DNA
yeast DNA
Apr
pMB1
ori
BgIII 5417
Tetr
YCp50
7.95 kb
URA3
NsiI 2488
XmaI 2540
SmaI 2542
BspMI 2845
ARS1
CEN4
SpeI 3294
991
HpaI 4099
34 Ks p 6311
37 Asu II
289 DraI
300 XhoI
184 GsuI
302 SciI
314 BgI II
339 Pvu II
351 BspHI
384 SpeI
413 BcII
577 ScaI
460 BcII
597 Avr II
630 SspI
776 Tth111I
914 DraI
922 DraI
948 DraI
1008 BspMI
1251 Eco 57I 1047 Avr II
1317 Asp718I
1191 HpaI
1321 KpnI
1357 GsuI
1402 StuI
1438 BsmI
1456 Ksp6321
1501 Eco 571 1648 BspHI
1524 Alw NI 1668 NheI
1534 DraI
1709 StuI
1746 Bs tXI
1776 Eco 31I
1792 XhoI
1794 SciI
2022 BaI I
2062 Eco 31I
2090 Eco RI
CEN4
885
XhoI 4702
BstXI 4649
KpnI 4227
NcoI 1866
AsuII 1926
ApaI 2039
Yeast Cloning
Vectors and Genes
Figure 13.4.5 YCp50. This vector is a derivative of YIp5 and YCp19. The EcoRI site of YCpl9 was
removed (producing an unsequenced deletion of about 190 bp) and a PvuII-HindIII fragment
(containing CEN4 and ARS1) from this derivative was cloned into the PvuII site of YIp5, with loss
of the PvuII site (Rose et al., 1987). Due to the presence of the CEN element, this plasmid exists
in low copy in yeast (1 to 2 copies/cell) and is mitotically stable (<1%loss per cell per generation).
This plasmid has not been completely sequenced; a more complete restriction map is available in
Rose et al., 1987.
A set of genomic plasmid banks using YCp50 and size-selected DNA fragments has been
constructed (Rose et al., 1987). These plasmid banks provide an alternative to genomic libraries
constructed in high-copy-number vectors, useful when isolating genes that would be lethal in yeast
when present in high copy.
Both the CEN3 (Fig. 13.4.7) and CEN4 (above) sequences were identified based on their ability
to confer mitotic stability and proper meiotic segregation to autonomously replicating plasmids
(Fitzgerald-Hayes et al., 1982; Mann and Davis, 1986). Nucleotide sequence comparison combined
with functional analysis has shown that centromeres contain three conserved structural elements.
Elements I and III show the highest degree of sequence conservation between different centromeres, and are separated by an extremely AT-rich region of about 90 bp, designated Element II.
Full CEN4 activity is contained within the 850-bp PvuII-HpaI fragment (which contains Elements I,
II, and III), although the adjacent 905-bp HpaI-EcoRI fragment also confers some mitotic stability
to unstable ARS-containing plasmids (Mann and Davis, 1986).
13.4.7
Supplement 30
Current Protocols in Molecular Biology
EcoRI
bacterial DNA
yeast DNA
EcoRI
TRP1
ARS1
CEN4
Apr
pMB1
ori
XhoI
HindIII
SUP4
ochre
pYAC3
11.4 kb
TEL
BamHI
SnaBI (pYAC3
cloning site)
BamHI
SaII
URA3
BamHI
HIS3
523
HIS3
DED1
827 HindIII
834 DraI
847 BgII
912 Bst XI
918 BgIII
821 Eco57I
978 BgI II
925 AsuII
935 SpII 1014 HindIII
1027 NheI
1071 BcII
1145 KpnI
1197 DraI/1206 PstI
1238 XcaI
1264 XcaI
1302 Eco57I/1302 NsiI
1331 BssHII
1379 XhoI
1381 SciI
1559 AfIII
1719 BgI II
1727 XmaIII
1750 CIaI
590 DraI
713 NdeI
730 BaII
735 BsmI
809 BvuII
475 AsuII
231 Eco 47III
49 NdeI/ 53 BamHI
105 DraI
114 Bvu II
PET56
BamHI
XhoI
HindIII
BamHI
HIS3
1179
TEL
Figure 13.4.6 pYAC3. pYAC vectors are used to clone very large fragments of exogenous DNA
onto artificial linear chromosomes, which can be stably maintained in yeast. This vector, which can
be propagated as a circular plasmid in E. coli, contains a unique cloning site in the SUP4 gene (an
ochre-suppressing allele of a tyrosine tRNA), as well as ARS1 and CEN4 elements, required for
stable single-copy propagation of the artificial chromosome. The TEL sequences are derived from
Tetrahymena telomeres and have been shown to function as telomeres in yeast. To clone an insert,
pYAC3 is digested with BamHI (which cuts adjacent to the telomere sequences) and SnaBI; the
resulting vector arms (containing either TRP1, ARS1, and CEN4 or URA3) are ligated to insert
fragments with SnaBI-compatible ends. The resulting ligation products are transformed into a ura3−
trp1− ade2-1 yeast strain, using the spheroplast protocol, selecting for Ura+ and subsequently
screening for Trp+ (to insure that both vector arms are present). Transformants can be further
screened for the presence of inserts in the middle of the SUP4 gene by using a color assay: colonies
in which the ade2-1 ochre mutation is suppressed by SUP4 are white, whereas inactivation of the
suppressor results in red colonies.
The pYAC vector shown above is one of a collection of three plasmids, each with a different
cloning site inserted into the SUP4 gene: pYAC4 and pYAC5 contain EcoRI and NotI sites,
respectively, in place of the SnaBI site found in pYAC3. Selected restriction sites (not necessarily
unique) are shown for pYAC3, as well as sites that have been destroyed in the process of plasmid
construction. The SUP4 gene is shown as a wavy line. For more detailed discussion of the cloning
protocol, as well as details of the construction of this vector, see Burke et al., 1987.
The HIS3 gene encodes imidazoleglycerolphosphate (IGP) dehydratase, which catalyzes a step
in the histidine biosynthetic pathway. This 1822-bp fragment also contains a portion of two other
genes: pet56, required for mitochondrial function, and ded1, required for cell viability (Struhl, 1985).
Mutations in the hisB gene of E. coli can be complemented by the cloned HIS3 yeast gene, using E.
coli BA1 (thr1 leuB6 trpC1117 hisB463 Tn10::near hisB thi1 thyA hsrk− hsmk− strr; Murray et al.,
1986).
Saccharomyces
cerevisiae
13.4.8
Current Protocols in Molecular Biology
Supplement 5
1753
Sal I
1987 SspI
2036 Bsp HI
2130 Pfl MI
2160 Asp 700I
2222 Tth 111I
1255 Ksp 632I
1295 Asp 700I/1295 EcoRI
1373 Bst XI
1408 EcoRV
1498 Hgi EII
1515 GsuI
1617 Eco NI
1635 Eco 57I
1652 Bsp MI
1708 Hgi EII
1840 Asp 700I
1110 Bbv II
780 AsuII
810 ClaI
209 Nar I
210 Eco 78I
212 BbeI
905 Asp718I
909 KpnI
927 Eco NI
692 Bst EII
486 Bsp HI
527 SspI
559 Pfl MI
241 HpaI
280 AsuII
3 Sci I
36 SspI
95 Ksp 632l
973 Afl II
LEU2
XhoI
95
647
LEU2
partial δ
100 bp
LYS2
NcoI
Pvu II
Hin dIII
100 bp
Eco RV
BamHI
XhoI
StuI
HpaI
NcoI
Eco RV
Bgl 2
EcoRI
LYS2 mRNA
Bam HI
574 Ksp 632I
591 Eco 57I
619 XmaI
621 SmaI
445 ScaI
100 bp
382 ScaI
395 ScaI
175
134 DraI
148 DraI
153 SspI
100 Dral
36 NheI
63
Sau 3A
CEN3
Figure 13.4.7 The LEU2 gene encodes β-isopropylmalate (β-IPM) dehydrogenase, which catalyzes the
third step in leucine biosynthesis (Andreadis et al., 1982). Unlike several other yeast genes involved in amino
acid biosynthesis, LEU2 (and LEU1, which is coordinately regulated with LEU2) is under specific amino acid
control: gene expression is repressed by elevated concentrations of leucine. The leu2-d allele is a deletion
of the 5′-flanking region of the LEU2 message which leaves only 29 bp preceding the LEU2 intiation codon;
this derivative of the LEU2 gene, when present on a YEp plasmid, requires a very high plasmid copy number
to give a Leu+ phenotype and has been used to cure cir+ strains of the endogenous 2µm plasmid (see UNIT
13.9). Also contained in this 2230-bp XhoI-SalI fragment are 95 bases of the 330-nucleotide δ element.
Although this δ element diverges in sequence from other δ elements, when the entire 2230-bp fragment is
used as a probe of genomic yeast DNA δ elements present elsewhere in the genome will be detected at a
low level. The cloned LEU2 gene can complement mutations in the leuB6 gene of E. coli using the strain
JA300 (thr1 leuB6 thi1 thyA trpC1117 hsrk− hsmk− strr; Tschumper and Carbon, 1982).
The LYS2 gene is the structural gene for α-aminoadipate reductase, which catalyzes an essential step
in lysine biosynthesis. The gene, which has not yet been sequenced, is present on a 4.6-kb EcoRI-HindIII
genomic fragment, and gives rise to a 4.2-kb LYS2 transcript, which is under general amino acid control
(Eibel and Philippsen, 1983; Barnes and Thorner, 1986). Much larger genomic fragments (up to 15.7 kb)
containing the LYS2 gene have been isolated, providing a large variety of restriction sites flanking the gene
for cloning purposes. As with URA3, a positive selection for lys2− mutants exists: such mutants can be
selected on medium containing α-aminoadipatic acid and lysine, with a spontaneous frequency of 10−5 to
10−6. Because the pathways for lysine biosynthesis in bacteria and fungi are not the same, no E. coli mutations
can be complemented by the cloned LYS2 gene.
See the legend to Figure 13.4.5 for a discussion of CEN3.
Yeast Cloning
Vectors and Genes
13.4.9
Supplement 5
Current Protocols in Molecular Biology
LITERATURE CITED
Andreadis, A., Hsu, Y.-P., Kohlhaw, G.B., and
Schimmel, P. 1982. Nucleotide sequence of yeast
LEU2 shows 5′ noncoding region has sequences
cognate to leucine. Cell 31:319-325.
Bach, M.L. LaCroute, F., and Botstein, D. 1979.
Evidence for transcriptional regulation of orotidine-5′-phosphate decarboxylase in yeast by hybridization of mRNA to the yeast structural gene
cloned in E. coli. Proc. Natl. Acad. Sci. U.S.A.
76:386-390.
Barnes, D.A. and Thorner, J. 1986. Genetic manipulation of Saccharomyces cerevisiae by use of the
LYS2 gene. Molec. Cell. Biol. 6:2828-2838.
Boeke, J., LaCroute, F., and Fink, G.R. 1984. A
positive selection for mutants lacking orotidine5′-phosphate decarboxlyase activity in yeast: 5fluoroorotic acid resistance. Mol. Gen. Genet.
197:345-346.
Botstein, D., Falco, S.C., Stewart, S.E., Brennan,
M., Scherer, S., Stinchcomb, D.T., Struhl, K.,
and Davis, R.W. 1979. Sterile host yeasts (SHY):
A eukaryotic system of biological containment
for recombinant DNA experiments. Gene 8:1724.
Burke, D.T., Carle, G.F., and Olson, M.V. 1987.
Cloning of large segments of exogenous DNA
into yeast by means of artificial chromosome
vectors. Science 236:806-812.
Clarke, L. and Carbon, J. 1978. Functional expression of cloned yeast DNA in Escherichia coli:
Specific complementation of argininosuccinate
lyase (argH) mutations. J. Mol. Biol. 120:517532.
Eibel, H. and Philippsen, P. 1983. Identification of
the cloned S. cerevisiae LYS2 gene by an integrative transformation approach. Mol. Gen. Genet.
191:66-73.
Elledge, S.J. and Davis, R.W. 1988. A family of
versatile centromeric vectors designed for use in
the sectoring-shuffle mutagenesis assay in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci.
U.S.A. In press.
Fitzgerald-Hayes, M., Clarke, L., and Carbon, J.
1982. Nucleotide sequence comparisons and
functional analysis of yeast centromere DNAs.
Cell 29:235-244.
Hartley, J.L. and Donelson, J.E. 1980. Nucleotide
sequence of the yeast plasmid. Nature 286:860865.
Hill, J.E., Myers, A.M., Koerner, T.J., and Tzagoloff,
A. 1986. Yeast/E. coli shuttle vectors with multiple unique restriction sites. Yeast 2:163-167.
Ma, H., Kunes, S., Schatz, P.J., and Botstein, D.
1987. Plasmid construction by homologous recombination in yeast. Gene 58:201-216.
Mann, C. and Davis, R.W. 1986. Structure and sequence of the centromeric DNA of chromosome
4 in Saccharomyces cerevisiae. Mol. Cell. Biol.
6:241-245.
Murray, A.W., Schultes, N.P., and Szostak, J.W.
1986. Chromosome length controls mitotic chromosome segregation in yeast. Cell 45:529-536.
Nasmyth, K.A. and Reed, S.I. 1980. Isolation of
genes by complementation in yeast: Molecular
cloning of a cell cycle gene. Proc. Natl. Acad.
Sci. U.S.A. 77:2119-2123.
Parent, S.A., Fenimore, C.M., and Bostian, K.A.
1985. Vector systems for the expression, analysis
and cloning of DNA sequences in S. cerevisiae.
Yeast 1:83-138.
Pouwels, P.H., Enger-Valk, B.E., and Brammar, W.J.
1985. Cloning Vectors: A Laboratory Manual.
Elsevier Science Publishing, Amsterdam.
Rose, M., Grisaffi, P., and Botstein, D. 1984. Structure and function of the yeast URA3 gene: expression in Escherichia coli. Gene 29:133-124.
Rose, M.D., Novick, P., Thomas, J.H., Botstein, D.,
and Fink, G.R. 1987. A Saccharomyces cerevisiae genomic plasmid bank based on a centromere-containing shuttle vector. Gene 60:237243.
Struhl, K. 1985. Nucleotide sequence and transcriptional mapping of the yeast pet56-his3-ded1
gene region. Nucl. Acids Res. 13:8587-8601.
Struhl, K., Stinchcomb, D.T., Scherer, S., and Davis,
R.W. 1979. High-frequency transformation of
yeast: Autonomous replication of hybrid DNA
molecules. Proc. Natl. Acad. Sci. U.S.A.
76:1035-1039.
Tschumper, G. and Carbon, J. 1980. Sequence of a
yeast DNA fragment containing a chromosomal
replicator and the TRP1 gene. Gene 10:157-166.
Contributed by Victoria Lundblad
University of California, Berkeley
Berkeley, California
Saccharomyces
cerevisiae
13.4.10
Current Protocols in Molecular Biology
Supplement 5
Yeast Vectors and Assays for Expression
of Cloned Genes
CONSTRUCTION OF LACZ FUSION VECTORS FOR STUDYING YEAST
GENE REGULATION
UNIT 13.6
BASIC
PROTOCOL 1
The lacZ gene of Escherichia coli encodes the enzyme β-galactosidase, which hydrolyzes
a variety of β-D-galactosides including chromogenic substrates (colorless compounds that
yield a colored product when hydrolyzed). Because of the ease and sensitivity of the
assays (see Basic Protocol 2 and see Alternate Protocol), yeast genes are often “tagged”
with a functional portion of the lacZ gene in order to monitor the regulation of expression
of the yeast gene in question. These fusions are constructed such that the promoter region
of the yeast gene—plus several amino acids from the N terminus of the protein encoded
by this gene—is fused to the carboxy-terminal region of the lacZ gene, which encodes a
protein fragment that still retains β-galactosidase activity. The liquid β-galactosidase
assay (see Basic Protocol 2) is sensitive and quantitative; it is used to accurately monitor
gene expression. The filter assay (see Alternate Protocol) is less sensitive and provides
only a qualitative assessment of β-galactosidase activity. However, the filter assay is
particularly useful for simultaneously and rapidly analyzing a large number of yeast
colonies.
When constructing lacZ fusions, it is crucial that the translational reading frame across
the fusion junction is maintained. If the sequence of the yeast gene is known, an in-frame
fusion can be engineered by choosing an appropriate fragment containing the promoter
region of the yeast gene as well as the N terminus of the encoded protein. Although the
same unique site preceding the lacZ fragment may not be present in the N terminus of the
yeast gene (and if present, may not generate an in-frame fusion), ligation of flush ends
(using another N-terminal restriction site) or the use of oligonucleotide linkers should
result in an in-frame fusion. If the sequence of the yeast gene is not known, in-frame
fusions can be identified empirically by assaying potential fusions for β-galactosidase
activity in either E. coli or yeast. A more detailed discussion of construction of in-frame
fusions can be found in Guarente (1983).
Figure 13.6.1 shows a vector, pLG670-Z, that can be used for constructing lacZ fusions
(Guarente, 1983). This plasmid contains the 2µm origin of replication and URA3 as a
selectable marker in yeast, with a unique BamHI site at the 5′ end of the lacZ fragment
(this fragment is actually itself a translational fusion of the lacI and lacZ genes, but for
simplicity will be referred to as the lacZ fragment). Preceding the BamHI site are unique
XhoI, Sa1I, and SmaI sites, which can aid in the insertion of various yeast fragments. The
XhoI and Sa1I sites are in a region of yeast DNA derived from the promoter region of the
CYC1 gene; however, a deletion of a portion of this promoter has inactivated it. The
translational reading frame of the lacZ gene immediately following the BamHI site is also
shown in Figure 13.6.1.
Two other plasmids (pLG200 and pLG400)—containing different translational reading
frames and/or different unique restriction sites at the 5′ end of the lacZ fragment—have
also been constructed (Guarente et al., 1980). These two plasmids do not contain yeast
selectable genes or yeast replication origins but can be used to first construct an in-frame
fusion, which is then transferred onto a yeast shuttle vector (UNIT 13.4).
Saccharomyces
cerevisiae
Contributed by Ann Reynolds, Victoria Lundblad, David Dorris, and Marie Keaveney
Current Protocols in Molecular Biology (1997) 13.6.1-13.6.6
Copyright © 1997 by John Wiley & Sons, Inc.
13.6.1
Supplement 39
bacterial DNA
yeast DNA
2µm origin
Ampr
URA3
pLG670-Z
XhoI
lacl
pMB1
ori
SmaI
SaII
lacZ
5′-GGA TCC GGA GCT TGG CTG-3′
BamHI
Figure 13.6.1 pLG670-Z (derived from YEp24) contains the 2µm origin of replication, URA3 as a
selectable yeast gene, and a 3′ fragment of the lacZ gene with a unique BamHI site at the 5′ end
of the lacZ fragment. The sequence just after the BamHI, as well as the translational reading frame,
is shown.
BASIC
PROTOCOL 2
ASSAY FOR â-GALACTOSIDASE IN LIQUID CULTURES
This protocol describes a rapid, quantitative assay of β-galactosidase activity in liquid
cultures of yeast. Yeast cells are either permeabilized or broken open, and the chromogenic
substrate o-nitrophenyl-β-D-galactoside (ONPG) is added in excess. After incubation at
30°C, the reaction is stopped by raising the pH to 11, inactivating β-galactosidase. Product
formation is determined spectrophotometrically.
Materials
YPD or other appropriate medium (UNIT 13.1)
Yeast strain containing lacZ fusion gene (see Basic Protocol 1)
Appropriate inducing agent (optional)
Z buffer (see recipe)
0.1% sodium dodecyl sulfate (SDS)
Chloroform
4 mg/ml ONPG (Table 1.4.2) in 0.1 M potassium phosphate, pH 7.0 (APPENDIX 2;
filter sterilized and stored frozen)
1 M Na2CO3
30°C water bath
Grow and prepare the cells
1. Inoculate 5 ml YPD (or appropriate) medium with a single yeast colony. Grow two
to three independent single-colony cultures of a yeast strain containing a lacZ fusion
gene overnight at 30°C.
Yeast Vectors for
Expression of
Cloned Genes
Assaying in duplicate or triplicate will help increase the accuracy. If the fusion is present
on a plasmid, cells should be grown in medium that selects for the plasmid.
13.6.2
Supplement 39
Current Protocols in Molecular Biology
2. Inoculate 5 ml YPD medium (or appropriate selective medium with or without
inducing agent) with 20 to 50 µl of each overnight culture. Grow to mid- or late-log
phase: 0.5–1 × 108 cells/ml (OD600 = 2.0) for rich medium or 2–5 × 107 cells/ml (OD600
= 0.5 to 1.0) for minimal medium.
If the goal is to investigate the activity of the yeast promoter under conditions that induce
its expression, cells should be grown in parallel under inducing and noninducing conditions.
3. Centrifuge cells 5 min at 1100 × g (2500 rpm in a tabletop centrifuge). Resuspend in
an equal volume of Z buffer and place on ice.
If the anticipated level of β-galactosidase activity is low, it may be necessary to concentrate
the cells. For cells demonstrating 100 to 1000 U of activity within 30 min to 4 hr,
concentration is not necessary.
4. Determine OD600 for each sample.
If cells are in mid-log phase, no dilution is necessary to obtain an accurate OD reading;
however, readings above 0.7 are inaccurate.
5. Set up the following two reaction tubes for each sample (1 ml each), with mixing:
(a) 100 µl cells with 900 µl Z buffer and (b) 50 µl cells with 950 µl Z buffer.
6. Add 1 drop of 0.1% SDS and 2 drops chloroform to each sample using a Pasteur
pipet. Vortex 10 to 15 sec and equilibrate 15 min in a 30°C water bath.
The addition of SDS and chloroform permeabilizes the cells. Alternatively, break cells open
using 0.5 g of acid-washed glass beads per sample (UNIT 13.12). Pellet the cell debris and
measure β-galactosidase activity and total protein concentration of the lysate. Enzyme
activity can then be normalized to protein concentration for calculating specific activity.
Assay for â-galactosidase
7. Add 0.2 ml of 4 mg/ml ONPG and vortex 5 sec. Place in a 30°C water bath and begin
timing.
8. When a medium-yellow color has developed, stop the reaction by adding 0.5 ml of
1 M Na2CO3 and note the time.
For accuracy, the OD420 should be 0.3 to 0.7. With practice, the amount of color correlating
with this OD reading can be recognized visually.
9. Centrifuge cells 5 min at 1100 × g. Determine OD420 and OD550 of the supernatant.
If the cell debris has been well-pelleted, the OD550—which measures light scattering by
cell debris—is usually zero and therefore is not necessary to read.
10. Calculate units with the following equation:
U=
1000 × [(OD 420 ) − (1.75 × OD 550 )]
(t ) × (v) × (OD 600 )
where t = time of reaction (min)
v = volume of culture used in assay (ml)
OD600 = cell density at the start of the assay
OD420 = combination of absorbance by o-nitrophenol
and light scattering by cell debris
OD550 = light scattering by cell debris.
Saccharomyces
cerevisiae
13.6.3
Current Protocols in Molecular Biology
Supplement 39
ALTERNATE
PROTOCOL
SCREENING FOR â-GALACTOSIDASE-EXPRESSING YEAST COLONIES
USING A FILTER LIFT ASSAY
This protocol (based on Durfee et al., 1993, and Staudinger et al., 1993) describes a
relatively simple and rapid method for simultaneously analyzing a large number of yeast
colonies for their ability to express β-galactosidase. This technique, based on a blue/white
color assay of colonies immobilized on filters, is much less laborious than the conventional liquid assay for β-galactosidase (see Basic Protocol 2). This assay can be used to
qualitatively measure the level of β-galactosidase expressed by individual yeast colonies
based on the intensity of the blue color, but it is less sensitive and quantitative than the
liquid β-galactosidase assay. The protocol is conveniently used as a secondary screen on
potential two-hybrid positive colonies (UNIT 20.1) that have already come through a
previous screening procedure or in genetic screens for mutations affecting the expression
of a specific reporter construct.
Materials
Yeast strain containing lacZ fusion gene (see Basic Protocol 1)
Selective medium plates (UNIT 13.1)
Liquid nitrogen
Z buffer (see recipe)
20 mg/ml Xgal (Table 1.4.2) in dimethylformamide
30°C incubator
Circular nitrocellulose membrane filters
Whatman 3MM paper
1. Spot or streak yeast colonies onto a plate containing appropriate selective medium
and allow to grow at 30°C until they reach optimal size (generally ∼48 hr).
Assaying colonies in duplicate will increase the reliability of the results.
2. Place a circular nitrocellulose membrane filter on the plate and press it gently on the
surface so that all the colonies are transferred to the membrane.
It is advisable to use reinforced nitrocellulose membranes for this procedure and treat them
very gently, as they become extremely fragile when placed in liquid nitrogen.
3. Carefully peel off the membrane and place in liquid nitrogen.
To avoid shattering of the membrane, use a shallow dish containing just enough liquid
nitrogen to cover the membrane.
4. Carefully remove the membrane from the liquid nitrogen container using an instrument such as a broad kitchen spatula. Place the membrane on a piece of dry Whatman
paper and allow to thaw (<5 min) at room temperature.
5. Place a circular piece of Whatman 3MM paper (cut to fit) into a petri plate. Add 3 to
5 ml of Z buffer containing 1 mg/ml Xgal—enough liquid to soak the paper without
flooding it. Place the thawed nitrocellulose membrane onto the Whatman paper and
allow the buffer to absorb slowly into the membrane.
Flooding the Whatman 3MM paper will allow diffusion of the colonies and make the
screening difficult to interpret.
6. Place the petri dish at 30°C and incubate a few minutes to overnight.
Strong positives should give a blue color after a few minutes of incubation time.
Yeast Vectors for
Expression of
Cloned Genes
13.6.4
Supplement 39
Current Protocols in Molecular Biology
REAGENTS AND SOLUTIONS
Z buffer
16.1 g Na2HPO4⋅7H2O (60 mM final)
5.5 g NaH2PO4⋅H2O (40 mM final)
0.75 g KC1 (10 mM final)
0.246 g MgSO4⋅7H2O (1 mM final)
2.7 ml 2-mercaptoethanol (50 mM final)
Adjust to pH 7.0 and bring to 1 liter with H2O. Do not autoclave.
COMMENTARY
Background Information
Cleavage of ONPG by β-galactosidase
yields two products, galactose and o-nitrophenol. The o-nitrophenol product is yellow and
can be detected by its absorption at 420 nm.
Because ONPG is present in excess in this
assay, the amount of o-nitrophenol produced is
proportional to the amount of enzyme present.
This assay is essentially that same as that used
for E. coli (Miller, 1972).
Critical Parameters
Liquid assay. Always use exponentially
growing cells and always normalize results by
measuring lacZ activity from a control strain
containing a previously characterized lacZ fusion protein that has been grown in parallel.
Reproducibility between measurements taken
on different days can be ensured by using the
same volume of inoculum in step 2 of Basic
Protocol 2 and growing cells to the same OD600
each time. The length of the reaction should be
from 30 min to 4 hr. Reaction times shorter or
longer than this will affect the accuracy of the
reaction. Adjust the reaction time by varying
the amount of Z buffer used to resuspend the
cells. If the fusion is present on a plasmid that
is lost at a high frequency (≥10%), the percentage of plasmid-bearing cells in the culture
should be determined (in step 2) and a correction made in calculating units of β-galactosidase activity.
LacZ filter lift assay. Always use both positive and negative control strains in this assay.
The positive control strain should give a strong
blue color after a few minutes of incubation
time. The negative control is especially important as this may turn a faint shade of blue
depending on the strain background and the
length of time of development.
Anticipated Results
Liquid assay. For a highly inducible fusion
(such as a GAL1 fusion) 5000 U can be observed, whereas other fusions (such as a HIS3lacZ fusion) have only 1 to 3 U. In some cases,
highly inducible fusions on high-copy-number
plasmids (such as 2 µm) may give values lower
than expected, presumably due to precipitation
of excess β-galactosidase present in the cell; to
increase assay accuracy, it may be necessary to
clone such fusions onto low-copy YCp vectors
or into the yeast genome.
LacZ filter lift assay. A strong two-hybrid
interaction strain should give a deep blue color
after just a few minutes of development time.
Weaker two-hybrid interaction strains may require up to several hours of incubation, and in
this instance they should be carefully compared
to the negative control to ensure that the color
change is not due to strain background.
Time Considerations
For the liquid assay, once mid-log cultures
are available, preparation of six samples will
take ∼45 min. The reaction time can vary between 30 min and 4 hr. Determining OD420 and
OD550 and calculating units require 15 to 30
min. Once the yeast plates are grown, the filter
lift assay takes 20 to 30 min followed by anywhere from 5 min to overnight for development, depending on the signal.
Literature Cited
Durfee, T., Becherer, K., Chen, P.-L., Yeh, S.-H.,
Yang, Y., Kilburn, A.-E., Lee, W.-H., and
Elledge, S.J. 1993. The retinoblastoma protein
associates with the protein phosphatase type 1
catalytic subunit. Genes & Dev. 7:555-569.
Guarente, L. 1983. Yeast promoters and lacZ fusions
designed to study expression of cloned genes in
yeast. Methods Enzymol. 101:181-191.
Saccharomyces
cerevisiae
13.6.5
Current Protocols in Molecular Biology
Supplement 39
Guarente, L., Lauer, G., Roberts, T.M., and Ptashne,
M. 1980. Improved methods for maximizing
expression of a cloned gene: A bacterium that
synthesizes rabbit β-globin. Cell 20:543-553.
Miller, J.H. 1972. Experiments in Molecular Genetics. Cold Spring Harbor Laboratory Press, Cold
Spring Harbor, N.Y.
Staudinger, J., Perry, M., Elledge, S.J., and Olson,
E.N. 1993. Interaction among vertebrate helixloop-helix proteins in yeast two-hybrid system.
J. Biol. Chem. 268:4608-4611.
Contributed by Ann Reynolds
University of Washington
Seattle, Washington
Victoria Lundblad
Baylor College of Medicine
Houston, Texas
David Dorris and Marie Keaveney
(filter lift assay)
Harvard Medical School
Boston, Massachusetts
Yeast Vectors for
Expression of
Cloned Genes
13.6.6
Supplement 39
Current Protocols in Molecular Biology
MANIPULATION OF YEAST GENES
SECTION III
Introduction of DNA into Yeast Cells
UNIT 13.7
The most commonly used yeast transformation protocol is the lithium acetate procedure
(basic protocol). It is reasonably fast and provides a transformation efficiency of 105 to
106 transformants/µg. This efficiency rivals that achieved for most, but not all, strains with
the more difficult and time-consuming spheroplast procedure (first alternate protocol).
However, the fastest and easiest of the transformation methods is electroporation (second
alternate protocol). For a number of strains, electroporation offers the highest transformation efficiency, and may prove especially useful with limiting quantities of transforming DNA. Unlike the lithium acetate procedure, however, electroporation saturates at low
DNA levels, restricting its general utility.
NOTE: All solutions and glassware coming into contact with yeast cells must be sterile.
Traces of soap on glassware may decrease the transformation efficiency. In addition, the
water used for washes and in solution preparation must be of the highest quality; see
reagents and solutions for guidelines.
TRANSFORMATION USING LITHIUM ACETATE
The lithium acetate method is based on the fact that alkali cations make yeast competent
to take up DNA. After yeast is briefly incubated in buffered lithium acetate, transforming
DNA is introduced with carrier DNA. Addition of polyethylene glycol (PEG) and a heat
shock trigger DNA uptake. The yeast are then plated on selective media.
BASIC
PROTOCOL
Materials
YPD medium (UNIT 13.1)
Yeast strain to be transformed
YPAD medium: YPD medium supplemented with 30 mg/liter adenine hemisulfate
Highest-quality sterile H2O
10× TE buffer, pH 7.5 (modify 1× recipe in APPENDIX 2), sterile
10× lithium acetate stock solution: 1 M lithium acetate, pH 7.5 (adjust pH
with dilute acetic acid), filter sterilized
DNA: high-molecular-weight, single-stranded carrier DNA (see support protocol)
and transforming DNA
50% (w/v) PEG 4000 or 3350 (do not use PEG 8000), filter sterilized
CM dropout plates (UNIT 13.1) prepared with Difco agar
30°C incubator with shaker
Sorvall GSA and SS-34 rotors (or equivalents)
42°C water bath
Grow and prepare the yeast cells
1. Two days before the experiment, inoculate 5 ml YPD medium with a single yeast
colony of the strain to be transformed. Grow overnight to saturation at 30°C.
The saturated overnight culture may, if desired, be prepared up to several weeks in advance
of the transformation and stored at 4°C.
2. The night before transformation, inoculate a 1-liter sterile flask containing 300 ml
YPAD medium with an appropriate amount of the saturated culture and grow
overnight at 30°C to 1 × 107 cells/ml (OD600 ≅ 0.3 to 0.5, depending on strain). For
2- to 3-fold higher efficiency, dilute at this point to 2 × 106 cells/ml in fresh YPAD
medium and grow for another 1 to 2 generations (2 to 4 hr).
Contributed by Daniel M. Becker and Victoria Lundblad
Current Protocols in Molecular Biology (1993) 13.7.1-13.7.10
Copyright © 2000 by John Wiley & Sons, Inc.
Saccharomyces
cerevisiae
13.7.1
Supplement 27
The presence of adenine in the medium produces a slightly higher transformation efficiency,
especially with ade− strains.
It is often difficult to know in advance how large an inoculum from the overnight culture
will produce a density of 1 × 107 cells/ml at a reasonable time the next day. Because growth
phase and cell density are important in achieving the highest transformation efficiency, the
easiest approach is to inoculate three independent flasks with varying amounts of the
saturated culture (try 1, 5, and 25 ìl for a 12-hr growth period), then check the OD600 of
each flask the next day.
3. Harvest cells by centrifuging 5 min at 4000 × g (5000 rpm in Sorvall GSA rotor),
room temperature. Resuspend in 10 ml highest-quality sterile water.
4. Transfer to a smaller centrifuge tube and pellet cells by centrifuging 5 min at 5000
to 6000 × g (7000 rpm in SS-34 rotor), room temperature.
5. Resuspend in 1.5 ml buffered lithium solution, freshly prepared as follows:
1 vol 10× TE buffer, pH 7.5
1 vol 10× lithium acetate stock solution
8 vol sterile water.
Cells may be incubated ≤1 hr at 30°C before adding the transforming DNA (step 6),
although this incubation is not required. In addition, the yeast may be stored ≤2 weeks at
4°C in this solution before proceeding with transformation. The transformation efficiency
will be much lower, but suffices for routine introduction of plasmids.
Transform yeast cells
6. For each transformation, mix 200 µg carrier DNA with ≤5 µg transforming DNA in
a sterile 1.5-ml microcentrifuge tube. Keep total volume of DNA ≤20 µl.
This protocol will yield a linear increase in the number of yeast transformants with
increasing input of transforming DNA over a large range (1 ng to 5 ìg).
Maximal transformation efficiency is achieved by repeating the denaturation cycle (boiling
and chilling) of carrier DNA immediately prior to use.
7. Add 200 µl yeast suspension to each microcentrifuge tube.
If the volume of DNA exceeds 20 ìl, add appropriate volumes of 10× lithium acetate stock
solution and 10× TE buffer to prevent further (>10%) dilution of the lithium acetate and
TE concentrations.
8. Add 1.2 ml PEG solution, freshly prepared as follows:
8 vol 50% PEG
1 vol 10× TE buffer, pH 7.5
1 vol 10× lithium acetate stock solution.
Shake 30 min at 30°C.
9. Heat shock exactly 15 min at 42°C. Microcentrifuge 5 sec at room temperature.
Longer or shorter duration of the heat shock will decrease transformation efficiency.
10. Resuspend yeast in 200 µl to 1 ml of 1× TE buffer (freshly prepared from 10× stock)
and spread up to 200 µl onto CM dropout plates made with Difco agar.
Difco agar gives 3-fold higher transformation efficiency than GIBCO/BRL agar.
11. Incubate at 30°C until transformants appear.
Introduction
of DNA into
Yeast Cells
Transformants will be visible on the surface of the agar 2 to 5 days after transformation,
depending on the strain, the plasmid, and the selection.
13.7.2
Supplement 27
Current Protocols in Molecular Biology
SPHEROPLAST TRANSFORMATION
This method is more time-consuming than the lithium procedure, but can result in a higher
efficiency of transformation per input DNA. Thus, it is the preferred method when the
DNA to be transformed is in limiting quantities or when a high efficiency of transformation is desirable—e.g., when screening a genomic plasmid bank.
ALTERNATE
PROTOCOL
Yeast cells are prepared for transformation by enzymatic digestion of the cell wall,
followed by several gentle isotonic washes (to remove the digestive enzyme). DNA is
mixed with the resulting spheroplasts and treated with a high-molecular-weight polymer,
polyethylene glycol (PEG), to promote DNA uptake. After removal of the PEG, spheroplasts and DNA are resuspended in osmotically stable regeneration agar and are plated
on appropriate selective plates.
Materials
YPD medium and plates (UNIT 13.1)
Yeast strain to be transformed
1 M sorbitol
2-mercaptoethanol (2-ME)
Glusulase (Du Pont NEN)
CaCl2 solution
Sorbitol/CaCl2 solution
DNA: transforming DNA and 5 mg/ml carrier DNA (sheared calf thymus or
salmon sperm DNA; UNIT 6.3)
PEG/CaCl2 solution
Selective regeneration agar
CM drop-out plates (UNIT 13.1)
30°C incubator with rotating platform
55°C water bath
NOTE: Before starting, melt selective regeneration agar (microwave at low setting),
aliquot 10-ml samples into sterile glass test tubes, and place in a 55°C water bath.
Grow the yeast strain
1. Two days before the experiment, inoculate 5 ml YPD medium with a single yeast
colony of the strain to be transformed. Grow overnight to saturation at 30°C (or lower
temperature, if strain is temperature sensitive).
2. The night before transformation, inoculate a 250-ml sterile flask containing 50 ml
YPD with an appropriate amount of the 5-ml culture and grow overnight to
1–2 × 107 cells/ml (OD600 ≅ 0.5 to 1.0, depending on strain).
The frequency of transformation drops with cell densities higher than this. If the exact
inoculum for a given strain is not known, it is best to inoculate three independent flasks
with varying amounts of the saturated 5-ml overnight as described in step 2 of the basic
protocol.
Prepare the spheroplasts
3. Pellet cells in a tabletop centrifuge 5 min at 1100 × g (2500 rpm) and resuspend in
10 ml of 1 M sorbitol. Repeat.
These and subsequent spins/resuspensions can be performed at room temperature.
4. Pellet cells once more and resuspend in 5 ml of 1 M sorbitol. Make 10−5 dilution in
sterile water and plate 0.1 ml onto a YPD plate. Incubate 2 days at 30°C.
5. Transfer resuspended cells to a 50-ml sterile flask and add 5 µl 2-ME and 150 µl
glusulase. Incubate ∼30 to 60 min at 30°C with very gentle shaking. Plate 0.1 ml of
Saccharomyces
cerevisiae
13.7.3
Current Protocols in Molecular Biology
Supplement 18
a 10−3 dilution (made in sterile water) onto a YPD plate and incubate 2 days at 30°C.
The number of colonies that grow up on this plate can be compared to the number that
grew on the plate from step 4 to calculate the percentage of spheroplast formation. Each
new lot of glusulase and each new strain should be assayed for the percentage of
spheroplast formation.
Spheroplast formation can also be monitored during the 30- to 60-min incubation with
glusulase by observing the degree of lysis in water under a microscope (spheroplasts will
lyse in a nonisotonic medium such as water, whereas intact yeast cells will not). At 15-min
intervals, remove 10 ìl of the cells, place on a microscope slide, and cover with a cover
slip. After focusing (at 240× to 400× magnification), gently touch a drop of water to the
edge of the cover slip. As the water leaks under the cover slip, observe how many cells lyse.
Spheroplasts should swell slightly before lysis; upon lysis, a “ghost” (membrane) should
still be visible.
6. Transfer spheroplasts to a sterile 50-ml, round-bottom centrifuge tube and pellet in
a tabletop centrifuge 4 min at 400 × g (1500 rpm). Gently decant supernatant without
dislodging pellet. Add 2 ml of 1 M sorbitol; resuspend pellet by gently swirling liquid
across surface of pellet (which should easily come off the side of the tube).
DO NOT vortex or otherwise vigorously agitate spheroplasts. If pellet does not easily
resuspend, reduce either the time or the rpm of subsequent spins.
7. Add 8 ml of 1 M sorbitol and pellet at 400 × g. Repeat steps 6 and 7.
8. Repeat step 6, add 7 ml of 1 M sorbitol and 1 ml CaCl2 solution, mix gently by swirling
and centrifuge 4 min at 400 × g.
9. Resuspend in 1 ml sorbitol/CaCl2 solution.
It has been reported that spheroplasts can be prepared and stored at −70°C in 1 M
sorbitol/15% DMSO and subsequently rethawed for use, with about a 5- to 10-fold
reduction in transformation efficiency (Orr-Weaver et al., 1983).
Transform the spheroplasts
10. Mix 150 to 200 µl cells with the DNA to be transformed (up to 10 µg) plus 10 µl
carrier DNA. Incubate 10 min at room temperature.
The total volume of added DNA should be no greater than 1⁄10 the volume of the cells. For
each transformation, it is advisable to mix an aliquot of cells with only carrier DNA, to
monitor the frequency of reversion of the relevant mutation.
The amount of transforming DNA to be added depends on several parameters, including
the frequency with which the strain can be transformed (determined experimentally) and
the state of the DNA (fragments of DNA without replication origins will transform several
orders of magnitude less efficiently than intact YEp, YRp or YCp plasmids—see UNIT 13.4).
11. Add a 10-fold volume of PEG/CaCl2 solution and thoroughly resuspend. Let sit 10
min at room temperature.
12. Pellet in tabletop centrifuge 4 min at 400 × g and decant the PEG-containing
supernatant.
Cells can also be plated in regeneration agar without removal of PEG, as long as the volume
of PEG + cells is 1⁄10 to 1⁄20 the volume of regeneration agar added. The presence of PEG,
however, will cause a decline in transformation frequency.
Introduction
of DNA into
Yeast Cells
13. Gently resuspend pellet in 0.5 ml of sorbitol/CaCl2 solution and pipet into 10 ml
melted regeneration agar tempered to 55°C. Vortex briefly to mix and immediately
pour onto the appropriate CM dropout (selective) plate, swirling the plate to further
mix cells and agar together.
13.7.4
Supplement 18
Current Protocols in Molecular Biology
The 3% agar will quickly solidify at temperatures <55°C, so it is important to plate each
10-ml aliquot as rapidly as possible.
14. Incubate at 30°C (or other appropriate temperature) until colonies appear, both on
the surface of the regeneration agar overlay and embedded in the agar. Pick individual
colonies and streak for single colonies on the same selective plates. If the purpose of
this transformation was to disrupt genomic sequences (UNIT 13.10), make DNA from
several transformants in order to analyze the relevant chromosomal region by
Southern blot analysis (UNIT 13.11).
TRANSFORMATION BY ELECTROPORATION
Yeast cells are concentrated 1000-fold from a log-phase culture using multiple washes
with sterile water to remove all extracellular ions. The concentrated cells are suspended
in 1 M sorbitol for osmotic stabilization. DNA is introduced without carrier, and the cells
are transformed using an exponential electric pulse delivered by an electroporation device.
The transformed cells are spread on a selective agar plate containing 1 M sorbitol. For
maximal efficiency, yeast may be incubated before concentration with DTT and lithium
acetate to render the cell wall/membrane more permeable to DNA.
ALTERNATE
PROTOCOL
Additional Materials
1 M dithiothreitol (DTT; filter sterilize and store at −20°C)
1 M sorbitol
Sorbitol selection plates
Gene Pulser with Pulse Controller (Bio-Rad) or Cell-Porator (GIBCO/BRL)
0.2-cm-gap disposable electroporation cuvettes (Bio-Rad) or 0.15-cm-gap
microelectroporation chambers (GIBCO/BRL); ice-cold
Grow and harvest the yeast cells
1. Two days before the experiment, inoculate 5 ml of YPD medium with a single yeast
colony of the strain to be transformed. Grow overnight to saturation at 30°C.
The saturated overnight culture may be prepared up to several weeks in advance of the
transformation and stored at 4°C.
2. The night before transformation, inoculate a 2-liter sterile flask containing 500 ml
YPD with an appropriate amount of the saturated culture and grow overnight
with vigorous shaking at 30°C to 1 × 108 cells/ml (OD600 ≅ 1.3 to 1.5, depending on
strain).
This cell density is achieved in mid- to late-log phase. It is often difficult to know how much
inoculum from the overnight culture will produce a cell density of 1 × 108 cells/ml at a
reasonable time the next day. If the exact inoculum for a given strain is not known, inoculate
three independent flasks with varying amounts of the saturated culture as described in step
2 of the basic protocol.
3. Harvest culture by centrifuging at 4000 × g (5000 rpm) 4°C, and resuspend vigorously
in 80 ml sterile H2O. To increase electrocompetence of the cells, proceed to step 4.
If this treatment is not required, proceed to step 7.
The rotor, the exact speed, and the duration of centrifuge spins are not critical; the principal
consideration is to centrifuge sufficiently hard (usually ∼4000 × g) to pellet all of the yeast,
but not so hard as to make resuspension difficult. For large volumes, 5000 rpm in a Sorvall
GSA rotor will suffice; for smaller volumes, 7000 rpm in a Sorvall SS-34 rotor is
recommended. This and subsequent centrifugation steps should be conducted at 4°C.
Saccharomyces
cerevisiae
13.7.5
Current Protocols in Molecular Biology
Supplement 18
Treat with lithium acetate and DTT to increase electrocompetence
This treatment is optional. It increases the handling time in preparing the yeast and should
not be performed if the cells are to be frozen for subsequent use.
4. Add 10 ml of 10× TE buffer, pH 7.5. Swirl to mix.
5. Add 10 ml of 10× lithium acetate stock solution (see first basic protocol). Swirl to
mix. Shake gently 45 min at 30°C.
6. Add 2.5 ml of 1 M DTT while swirling. Shake gently 15 min at 30°C.
This treatment increases the transformation efficiency >5-fold. Both lithium acetate and
DTT, if used individually, will increase transformation efficiency. Used together, the
improvement is approximately additive.
Concentrate and wash the yeast cells
7. Dilute yeast suspension to 500 ml with water.
8. Wash and concentrate the cells three times by centrifuging at 4000 to 6000 × g,
resuspending the successive pellets as follows:
First pellet—250 ml ice-cold water
Second pellet—20 to 30 ml ice-cold 1 M sorbitol
Third pellet—0.5 ml ice-cold 1 M sorbitol.
Resuspension should be vigorous enough to completely dissociate each pellet. The final
volume of resuspended yeast should be 1.0 to 1.5 ml and the final OD600 should be ∼200.
Yeast can be stored at −70°C for subsequent use by addition of glycerol to 15% (v/v)
followed by freezing in a dry ice/ethanol bath. Frozen aliquots should be thawed slowly,
pelleted in a sterile microcentrifuge tube, and resuspended to the same volume in 1 M
sorbitol. The wash is necessary to remove ions liberated by lysis of some yeast cells.
Transformation efficiency will drop ≥10-fold with freezing.
Electroporate the yeast cells
Using the Bio-Rad Gene Pulser:
9a. In a sterile, ice-cold 1.5-ml microcentrifuge tube, mix 40 µl concentrated yeast cells
with ≤100 ng transforming DNA contained in ≤5 µl.
Transforming DNA should be in a low-ionic-strength buffer such as TE or in sterile
high-quality water (see reagents and solutions). There is no required length of incubation;
this time can be varied to convenience. Do not include carrier DNA in this procedure; it
drastically reduces transformation efficiency. Maximal efficiency (transformants/ìg) will
be obtained with <10 ng of transforming DNA, while the largest number of transformants
will be obtained with 100 ng. Using >100 ng will cause a precipitous drop in both
transformation efficiency and the total number of transformants, with few colonies evident
in transformation with 1 ìg DNA.
10a. Transfer to an ice-cold 0.2-cm-gap disposable electroporation cuvette.
11a. Pulse at 1.5 kV, 25 µF, 200 Ω. It is crucial that the Bio-Rad Pulse Controller be
included in the circuit; failure to do so will result in damage to the Gene Pulser.
The time constant reported by the Gene Pulser will vary from 4.2 to 4.9 msec. Times <4
msec or the presence of a current arc (evidenced by a spark and smoke) indicate that the
conductance of the yeast/DNA mixture is too high.
Introduction
of DNA into
Yeast Cells
12a. Add 1 ml ice-cold 1 M sorbitol to the cuvette and recover the yeast, with gentle
mixing, using a sterile 9-inch Pasteur pipet.
13.7.6
Supplement 18
Current Protocols in Molecular Biology
13a. Spread aliquots of the yeast suspension directly on sorbitol selection plates. Incubate
at 30°C until colonies appear.
Although there is a conflicting report (Lorow-Murray and Jessee, 1991), we find that the
addition of sorbitol as an osmotic protectant to the agar increases survival of transformed
yeast, yielding 10-fold higher transformation efficiency (Becker and Guarente, 1991).
Transformants will be visible on the agar surface 3 to 6 days after transformation,
depending on the strain, the plasmid, and the selection. This is usually 1 day longer than
with the lithium acetate procedure.
Using the BRL Cell-Porator:
9b. In a sterile, ice-cold 1.5-ml microcentrifuge tube, mix 20 µl concentrated yeast with
≤100 ng transforming DNA contained in ≤5 µl.
Incubation time can be varied for convenience (step 9a).
10b. Transfer to an ice-cold, 0.15-cm-gap micro-electroporation chamber.
11b. Pulse at 400V, 10 µF, low resistance.
12b. Remove 10 µl of electroporated mixture to a sterile 1.5-ml microcentrifuge tube
containing 0.5 ml ice-cold 1 M sorbitol.
13b. Spread aliquots of the yeast suspension directly on sorbitol selection plates. Incubate
at 30°C until colonies appear.
PREPARATION OF SINGLE-STRANDED HIGH-MOLECULARWEIGHT CARRIER DNA
SUPPORT
PROTOCOL
Addition of denatured high-molecular-weight carrier DNA is critical to achieving high
transformation efficiency using the lithium acetate protocol. Both parameters are crucial:
the DNA must be single-stranded, and the larger the fragments, the better the transformation efficiency. Carrier DNA prepared according to this support protocol may also be used
for spheroplast transformations, although single-stranded DNA has been reported to give
a slightly lower transformation frequency than the more easily prepared double-stranded
calf thymus or salmon sperm carrier DNA. (NOTE: Addition of this or any other carrier
DNA during electroporation drastically reduces the efficiency of transformation.)
Materials
DNA (type III sodium salt from salmon testes; Sigma #D1626)
1× TE buffer, pH 8.0 (APPENDIX 2)
Buffered phenol (UNIT 2.1)
1:1 (v/v) phenol/chloroform
Chloroform
3 M sodium acetate, pH 5.2 (APPENDIX 2)
100% ethanol, ice-cold
Probe sonicator
Additional reagents and equipment for agarose gel electrophoresis (UNIT 2.5) and
phenol extraction and ethanol precipitation of DNA (UNIT 2.1)
1. Dissolve DNA in 1× TE buffer, pH 8.0, to a final concentration of 10 mg/ml. Stir
overnight at 4°C.
The solution of DNA will be extremely viscous.
Because the correct preparation of carrier DNA is crucial in achieving high transformation
efficiency by the lithium acetate protocol, and because each individual transformation
Saccharomyces
cerevisiae
13.7.7
Current Protocols in Molecular Biology
Supplement 18
requires 200 ìg, prepare a large amount (e.g., 1 g) and freeze aliquots at −20°C.
2. Sonicate using a large probe until viscosity appears to decrease slightly. Run 1 µg on
a 0.8% agarose gel to determine size distribution of sonicated fragments. Repeat
sonication as necessary to achieve the appropriate size distribution.
Each sonication should be brief— ∼30 sec at 75% power (see UNIT 6.3 for guidelines on
sonication of herring sperm DNA). The larger the fragments, the better the transformation
efficiency will be, but the more viscous and unwieldy the solution. The optimum distribution
of fragment size is that in which fragments range from 2 kb to 15 kb, with a mean size of
∼7 kb.
3. Extract once with buffered phenol, once with phenol/chloroform, and once with
chloroform.
4. Precipitate DNA with 1⁄10 vol of 3 M sodium acetate, pH 5.2, and 2.5 vol ice-cold
100% ethanol.
With DNA of this size and at this concentration, the DNA will form flocculent masses
immediately upon addition of the ethanol.
5. Resuspend pellet with 1× TE buffer to 10 mg/ml final concentration.
Resuspension may require stirring overnight at 4°C.
6. Transfer carrier DNA to a Pyrex flask. Microwave to a rolling boil and continue to
boil 2 to 3 min.
7. Chill flask rapidly in ice water. Aliquot and freeze DNA in sterile tubes at −20°C.
REAGENTS AND SOLUTIONS
CaCl2 solution
0.1 M Tris⋅Cl, pH 7.4 (APPENDIX 2)
0.1 M CaCl2
Autoclave or filter sterilize and store at room temperature
PEG/CaCl2 solution
45% (w/v) PEG 3350
10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2)
10 mM CaCl2
Filter sterilize and store at room temperature
Selective regeneration agar
5 g ammonium sulfate
1.7 g yeast nitrogen base (without amino acids or ammonium sulfate)
20 g dextrose
30 g agar
1 pellet NaOH
1.3 g appropriate amino acid dropout powder (Table 13.1.1)
To above ingredients, add 500 ml of 2 M sorbitol, 20 ml YPD medium (UNIT 13.1),
and 480 ml water. Mix well and autoclave 15 min in 2-liter flask. Aliquot 250-ml
portions into 500-ml sterile bottles, using sterile technique. Store at room temperature or 4°C.
Introduction
of DNA into
Yeast Cells
Sorbitol selection plates
Supplement CM dropout plates (UNIT 13.1) with sorbitol to 1 M final concentration.
Sorbitol can be added as a powder prior to autoclaving. NOTE: Addition of sorbitol
makes the agar more viscous and more prone to boiling over upon removal from the
autoclave. Store at room temperature or 4°C.
13.7.8
Supplement 18
Current Protocols in Molecular Biology
Sorbitol/CaCl2 solution
1 M sorbitol
10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2)
10 mM CaCl2
Filter sterilize or autoclave and store at room temperature
Water, high-quality and sterile
Water used in these protocols for washes and in solution preparation must be of the
highest-possible quality (e.g., Millipore Milli-Q); the resistance must be at least as
high as 10 MΩ/cm. Sterilize by autoclaving and store at room temperature.
COMMENTARY
Background Information
High-frequency transformation in yeast was
first demonstrated using β-glucanase to remove
the cell wall to generate spheroplasts (Beggs,
1978; Hinnen et al., 1978). The method has
several major disadvantages. Of the procedures
for transforming yeast, it is the most tedious
and difficult. Yeast transformed by spheroplasting must be embedded in top agar to allow
regeneration of the cell wall, which makes
subsequent manipulations, such as replica plating or colony hybridization (UNIT 13.2), extremely difficult. In addition, the PEG treatment of spheroplasts often generates diploids
and triploids by cell fusion. Nonetheless,
spheroplast transformation has long remained
a preferred approach for yeast transformation
because it offered, until recently, the highest
transformation efficiencies for most yeast
strains.
More recent transformation protocols, such
as that employing lithium acetate (Ito et al.,
1983), do not require digestion of the cell wall
and thus do not require regeneration in top
agar—rather, transformants are plated on the
agar surface. With lithium acetate transformation, competence for DNA uptake at the level
of the cell wall is induced by treatment with
lithium ions plus PEG. This procedure is easier
and more rapid than spheroplasting. The finding that single-stranded, high-molecularweight carrier DNA could increase transformation efficiency by one to two orders of magnitude (Schiestl and Gietz, 1989) has made
lithium acetate transformation at least as efficient as spheroplast transformation. This modified lithium acetate procedure will likely replace spheroplast transformation for most
strains.
The final and simplest approach is electroporation (Becker and Guarente, 1991). Yeast
are harvested during log-phase growth, concentrated, and suspended in an osmotically stabi-
lizing medium with transforming DNA. A
high-voltage electrical discharge through this
suspension drives the DNA uptake. Electroporation can be extremely efficient, especially
with small quantities of DNA (1 pg to 10 ng).
When DNA quantities are limited, this may be
the procedure of choice. Electroporation has
several drawbacks, however. First, the expense
of the electroporator apparatus may be prohibitive for some labs. Second, the efficiency of
transformation is extremely strain-dependent.
Third, the process saturates; increasing the input above 100 ng decreases the yield of transformants. Electroporation is inappropriate,
therefore, for experiments that require maximizing the total number of transformants, such
as screening genomic or cDNA expression libraries in yeast.
Critical Parameters
Transformation efficiency in all of these
procedures depends on the yeast strain background. Lithium acetate transformation is least
affected by strain variation, while electroporation is most affected. In the spheroplast method,
strain-dependent variations in frequency may
reflect differing sensitivities to glusulase. In
electroporation, strain-dependent variations
may reflect differing degrees of survival after
the electric pulse. Since the response of a strain
cannot be predicted a priori, strains known to
transform at high efficiency should be used
whenever possible. When strain selection is
dictated by the particular experiment, all the
protocols might have to be tested to determine
which works best.
Maximal transformation efficiency in each
of the procedures also depends on finding the
optimal cell density. Each protocol lists the
preferred starting density; strain differences
may require empiric variation. The condition
of the DNA has an effect as well; miniprep
DNA prepared by either alkaline lysis or boil-
Saccharomyces
cerevisiae
13.7.9
Current Protocols in Molecular Biology
Supplement 18
ing (UNIT 1.6) will transform adequately, but
phenol extraction (UNIT 2.1) or purification by
CsCl/ethidium bromide density gradients (UNIT
1.7) will further increase the efficiency.
In the lithium acetate protocol, the single
most critical parameter is the carrier DNA. It
must be of high-molecular-weight and completely denatured (Schiestl and Gietz, 1989).
Denaturation can be assured by repeating the
cycle of boiling and chilling immediately before use. Achieving the proper size distribution
may be more difficult. If low efficiencies are
obtained with several strains, another batch of
carrier DNA should be prepared. In addition,
using Difco agar to prepare the CM dropout
plates increases the transformation efficiency.
In the spheroplast procedure, the most critical parameter is the handling of spheroplasts—
vortexing, osmotic shock, or plating in regeneration agar at temperatures >55°C will reduce
transformation frequency. For both the lithium
and spheroplast protocols, transformation efficiency also depends on the batch of PEG used.
If poor frequencies are obtained, several lots of
PEG must be tested. Be certain to use PEG
3350, not PEG 8000.
In electroporation, the most important parameter after strain selection is the amount of
transforming DNA used. Maximal efficiency
(transformants/µg) will be obtained with <10
ng of transforming DNA; the largest number of
transformants will be obtained with 100 ng.
Using more than 100 ng of plasmid or including
carrier DNA will cause a precipitous drop in
both transformation efficiency and total number of transformants. The electrical parameters
are also important. With either of the exponential decay devices described, strain differences
may require empiric variation of the initial
voltage around the suggested value. For squarewave devices, see Meilhoc et al. (1990).
For all protocols, it is important that sterile
technique be used throughout. Traces of soap
on glassware may decrease the transformation
efficiency.
Anticipated Results
Under optimal conditions, a circular plasmid (e.g., YCp or YEp; UNIT 13.4) should give
105 to 106 transformants/µg DNA with the
lithium acetate protocol, ∼104 to 105 by spheroplasting, and >105 with electroporation (>104
transformants at 100 ng). With either linear
molecules or circular molecules that do not
contain a yeast replication origin, stable transformation requires integration into the genome;
transformation frequencies are correspondingly lower, with ∼102 to 103 transformants/µg
DNA.
Time Considerations
Once the cells have reached the desired density, the lithium acetate protocol requires ∼3 hr
to complete, spheroplasting requires 3 to 4 hr,
and electroporation can be completed in ∼90
min. Because spheroplasts are metabolically
active for up to 24 hr, the time spent washing
the spheroplasts can be increased for convenience (e.g., each resuspension step can be left
for an hour or more on a gently rocking platform).
Literature Cited
Becker, D.M. and Guarente, L. 1991. High-efficiency transformation of yeast by electroporation. Methods Enzymol. 194:182-187.
Beggs, J.D. 1978. Transformation of yeast by a
replicating hybrid plasmid. Nature (Lond.)
275:104-109.
Hinnen, A., Hicks, J.B., and Fink, G.R. 1978. Transformation of yeast. Proc. Natl. Acad. Sci. U.S.A.
75:1929-1933.
Ito, H., Fukuda, Y., Murata, K., and Kimura, A. 1983.
Transformation of intact yeast cells treated with
alkali cations. J. Bacteriol. 153:163-168.
Lorow-Murray, D. and Jessee, J. 1991. Highefficiency transformation of Saccharomyces
cerevisiae by electroporation. Focus 13:65-67.
Meilhoc, E., Masson, J.-M., and Teissie, J. 1990.
High efficiency transformation of intact yeast
cells by electric field pulses. Bio/Technology
8:223−227.
Orr-Weaver, T.L., Szostak, J.W., and Rothstein, R.J.
1983. Genetic applications of yeast transformation with linear and gapped plasmids. Methods
Enzymol. 101:228-244.
Schiestl, R.H. and Gietz, R.D. 1989. High efficiency
transformation of intact yeast cells using single
stranded nucleic acids as a carrier. Curr. Genet.
16:339-346.
Contributed by Daniel M. Becker
Stanford Law School
Stanford, California
Victoria Lundblad
Baylor College of Medicine
Houston, Texas
Introduction
of DNA into
Yeast Cells
13.7.10
Supplement 18
Current Protocols in Molecular Biology
Cloning Yeast Genes by Complementation
Yeast genes have been cloned by a variety of techniques, including use of purified RNA
as hybridization probes, differential hybridization of regulated RNA transcripts, antibody
screening, transposon mutagenesis, cross suppression of mutant phenotypes, cross hybridization with heterologous cDNA or oligonucleotide probes, as well as by complementation in E. coli (for reviews of these methods see Rothstein, 1985, or Rose, 1987).
This unit presents a generalized protocol and describes the principles involved in cloning
yeast genes by complementation in yeast.
UNIT 13.8
BASIC
PROTOCOL
The protocol is presented using a hypothetical mutation of yeast, the cdc101-1 mutation.
This mutation was isolated as a cell cycle mutant and is both recessive and temperaturesensitive for growth: it can grow relatively normally at 30°C but is unable to make a colony
at 37°C. A genomic DNA clone that complements this mutation will be isolated by
transforming the cdc101-1 strain with a yeast genomic library and subsequently screening
for temperature-resistant colonies. Once isolated, two steps are necessary to prove that
the insert present on the plasmid contains the wild-type CDC101 gene. First, segregation
of the complementing plasmid must result in co-loss of both the plasmid-borne selectable
marker and the complementing phenotype, demonstrating that the observed complementation is plasmid-specific and is not due to reversion of the cdc101-1 mutation. Second,
it must be ruled out whether the cloned gene encodes a phenotypic suppressor of the
mutation, rather than the wild-type gene. This is done via a complementation test, which
demonstrates whether or not a disruption of the cloned gene that is integrated into the
genome can complement the original mutation.
Cloning the Gene
Transform a mutant yeast strain
1. Transform a leu2− cdc101-1 yeast strain with a yeast genomic DNA library containing
LEU2 as a selectable marker (using either protocol in UNIT 13.7) and selecting, in this
case, for Leu+. Depending on the insert size, between 2,000 and 20,000 transformants
must be screened (∼4 to 8 genome equivalents).
If the mutation reverts at a very low frequency (<10−8), transformants that complement the
mutant phenotype (in this case, temperature sensitivity) can be selected directly after
transformation. However, if the mutation reverts at higher frequencies (or if the complementing phenotype cannot be directly selected), the mutation must be present in a strain
that also contains a low or nonreverting mutation in the selectable gene present on the
library vector.
Whenever possible, it is also preferable to have the mutation present in a strain that
transforms well. If the mutation has been isolated in a strain that transforms poorly,
alternate transformation protocols can be tested (see UNIT 13.7). Transformation proficiency
can also be improved by backcrosses with a known strain that is more competent for
transformation.
Screen for complementation of the mutant phenotype
2. Replica plate transformants onto prewarmed selective plates (leucine dropout plates;
UNIT 13.1) and incubate at 37°C, to screen for complementation of the temperaturesensitive phenotype. With overnight incubation, colonies containing plasmids with
inserts that complement the cdc101-1 mutation, as well as potential CDC101+
revertants that contain random noncomplementing plasmids, will grow up. Restreak
each single colony that appears on this plate and save for further analysis.
Saccharomyces
cerevisiae
Contributed by Victoria Lundblad
Current Protocols in Molecular Biology (1989) 13.8.1-13.8.4
Copyright © 2000 by John Wiley & Sons, Inc.
13.8.1
Supplement 5
Many mutant phenotypes, including ts phenotypes, cannot be distinguished by replica
plating (presumably due to leakiness of the mutation). If this is the case, transformants
must be recovered from the original transformation plate and replated directly on a second
plate that screens for the mutant phenotype. If the lithium method of transformation is used,
recover transformants by pipetting about 0.5 ml sterile water onto each plate and resuspending the individual colonies in the liquid by mushing about the plate with a sterile
spreader (UNIT 1.3). Pipet the 0.5 ml back into a sterile tube and replate for single colonies
onto the appropriate plate. If the spheroplast protocol is used, chop up the agar from each
transformant plate (using a sterile spatula) and resuspend in 5 to 10 ml sterile water. Vortex
extensively to further break up the agar (which releases the transformants into the liquid)
and replate for single colonies.
If the mutation displays more than one phenotype, complementation of the additional
phenotypes should also be tested. Complementation of some but not all of the mutant
phenotypes indicates that the wild-type gene has not been cloned. However, such a
complementation pattern suggests that a gene which performs a related function has been
isolated.
Proof That the Correct Gene Has Been Cloned
Determine whether segregation of the complementing plasmid results in co-loss of
both the plasmid-borne selectable marker and the complementing phenotype
3. Using the protocol described in UNIT 13.9, isolate Leu− segregants for each candidate
and test their ability to grow at 37°C. Save transformants that are only able to grow
at 37°C in the presence of the plasmid.
If plasmid segregants can be isolated that still display “complementation,” this suggests
that the complementation pattern is due to either reversion of the original mutation or
acquisition of a suppressor mutation. Alternatively, if colonies are observed that still retain
plasmid but are no longer complemented for the mutant phenotype, this indicates that the
original transformant contained a mixed population of plasmids.
4. Isolate plasmid DNA from each transformant identified in step 3 by transforming
total yeast DNA into E. coli (UNIT 13.11). Retransform each plasmid into the mutant
yeast strain and confirm that the correct plasmid has been isolated (in our example,
demonstrate that retransformation into a cdc101-1 strain results in a temperature-resistant phenotype).
5. Analyze the plasmids by restriction mapping (UNIT 3.2) to determine whether genomic
DNA inserts present in different plasmids have overlapping segments, which can help
define the boundaries of the complementing region. Construction of various deletion
and insertion mutations, and tests of their ability to complement the mutation in yeast,
will provide more precise information about the location of the gene within the cloned
insert.
Determine whether a disruption of the cloned gene that is integrated into the
genome can complement the original mutation (complementation test)
The following steps are designed to test whether the cloned gene encodes a phenotypic
suppressor of the mutation, rather than the wild-type gene.
6. Introduce a yeast selectable marker (UNIT 13.4) into a site in the middle of the
complementing region (based on the information gained in step 5).
Cloning Yeast
Genes by
Complementation
7. Transform this disrupted plasmid back into the original mutant strain and screen
for the mutant phenotype to test whether complementation has been abolished as
follows. Using one of several techniques discussed in UNIT 13.10, construct a diploid
strain containing one wild-type copy of the gene and one disrupted copy of the
gene. After sporulation and dissection (UNIT 13.2), cross a haploid spore product
13.8.2
Supplement 5
Current Protocols in Molecular Biology
containing the disruption (usually identified by the presence of the selectable marker
associated with the disruption) is crossed to a strain carrying the original mutation
(UNIT 13.2). If this diploid has the same mutant phenotype as the strain with the original
mutation, this demonstrates lack of complementation and indicates that the wild-type
gene corresponding to this mutation has been cloned. As a control, cross the strain
with the gene disruption (which may itself display a non–wild-type phenotype) to a
wild-type strain, in order to demonstrate that the phenotype of this diploid is wild-type
(showing that the disruption is a recessive mutation).
Often, a genomic disruption of the cloned insert will have a mutant phenotype which is
either similar to that of the original mutation or is a phenotype predicted for a null mutation
in this gene. Although this is suggestive that the correct gene has been cloned, it does not
constitute proof: a related gene may not only compensate for the original mutation when
present on a plasmid, but may also give a similar phenotype when disrupted. Complementation, however, will demonstrate whether two different genes have been identified.
In the case of the hypothetical CDC101 gene, the cdc101-1 mutation is a ts lethal,
indicating that this is an essential gene and that a haploid strain containing a disruption
of the CDC101 gene would be inviable (techniques for determining whether this hypothesis
is correct are presented in UNIT 13.10). For such an essential gene, the same complementation test described above can be performed, with one technical modification. Construct a
plasmid-borne disruption of the insert region that complements cdc101-1, using URA3 as
the selectable marker. Transform a ura3−/ura3−, leu2−/leu2−, diploid strain with this
disruption (using techniques described in UNIT 13.10), selecting for Ura+. Now transform
this heterozygous diploid with a LEU2 plasmid bearing an intact copy of the complementing region. After sporulation, identify a spore product that contains both the URA3
disruption and the LEU2 complementing plasmid. This strain can now be crossed to both
a cdc101-1 strain and a control CDC101+ strain, the Leu+ plasmid segregated away (UNIT
13.9), and complementation (or lack of it) observed.
COMMENTARY
Cloning a gene by complementation in yeast
requires not only a low-reverting recessive mutation in the gene of interest, but the appropriate
yeast genomic DNA library. A number of yeast
genomic libraries have been published and may
be available (e.g., see Rose et al., 1987; MeeksWagner et al., 1986; Kuo and Campbell, 1983).
The parameters to consider when selecting a
preexisting library are the auxotrophic mutation(s) present in the strain of interest (which
determines the selectable marker that must be
present on the vector), whether the strain is cir+
or cir0 (libraries constructed in YEp plasmids
are much less stable in cir0 strains; see UNIT 13.4),
and the copy number of the vector (some genes,
when present in high copy, are toxic to yeast).
In some cases, it may be necessary to construct
a yeast plasmid library de novo. This is the case
when the only mutation available in yeast is
dominant to the wild-type allele. Construction
of a library from DNA isolated from the mutant
strain will allow isolation of the mutant gene
(by screening transformants of a wild-type
strain for the mutant phenotype), from which
the wild-type gene can be obtained by standard
cloning techniques (UNITS 6.1 to 6.3). In addition,
some yeast genes have been demonstrated to be
toxic when expressed in E. coli. Thus, preexisting yeast libraries which have been amplified
through E. coli will be under-represented for
plasmids containing these genes. This can be
solved by constructing a library and transforming directly into yeast. A more detailed discussion of the issues involved in choosing a library,
as well as a protocol for constructing a yeast
genomic library, are presented in Rose, 1987
(see also Chapter 5).
Cloning yeast genes by complementation
when the only available mutation is dominant
presents special problems. As mentioned
above, a genomic library must be constructed
from the mutant strain. In addition, once a
candidate gene has been isolated, if either the
original mutation or a genomic disruption in
this candidate gene are dominant, a complementation test cannot be used to prove that the
correct gene has been cloned. In this case,
another genetic test can be used to demonstrate
tight genetic linkage between a selectable
marker introduced at the chromosomal site of
the cloned gene and the original mutation. In
this test, a plasmid carrying the cloned gene as
Saccharomyces
cerevisiae
13.8.3
Current Protocols in Molecular Biology
Supplement 7
well as a selectable marker are integrated into
the genome of a wild-type haploid strain, using
the technique of integrative transformation
(UNIT 13.10). This haploid strain is crossed with
a strain carrying the (dominant) mutation, and
the subsequent diploid is sporulated and analyzed for the pattern of segregation of the mutation and the selectable marker associated with
the integrated gene. If the cloned gene has
integrated at the genetic locus corresponding to
the original mutation, the selectable marker
will always cosegregate with wild-type—i.e.,
spore products will never be recovered that
display both the mutation and the selectable
marker. If the cloned gene has integrated at
another unlinked genetic locus, the selectable
marker will segregate independently of the mutant/wild-type phenotypes, with approximately
one-fourth of the spores containing both the
mutation and the selectable marker (e.g., however, <25% of the spores will contain both
markers if the cloned DNA has integrated at a
site that is somewhat linked to the mutation). If
tight genetic linkage can be demonstrated in
this test, this provides evidence for the identity
of the cloned piece of DNA. However, it is
possible that a suppressor gene has been cloned
that is tightly linked to the original mutant
allele; such a possibility would not be ruled out
by the above test.
Cloning of heterologous genes by complementation in yeast follows the same general
principles presented in this section but requires
a library of genomic DNA or cDNA from the
relevant organism cloned into a vector that can
be maintained in yeast. Although in some cases
the heterologous gene can be expressed in yeast
by its own promoter, in most cases the gene
must be fused to a yeast promoter. Several
heterologous libraries containing DNA from
S. pombe (Beach et al., 1982) and Drosophila
have already been constructed. Vectors for ex-
pression of genes in yeast that can be used for
construction of heterologous libraries are discussed in UNIT 13.4. One principle that cannot be
applied to cloned heterologous genes is the use
of complementation (or linkage analysis, as
described in the previous paragraph) to prove
that the correct gene has been cloned. In these
situations, evidence that the correct gene has
been cloned may require comparison of nucleic
acid and protein sequence between the two
genes, as well as functional analysis (e.g., demonstrating that the two genes encode the same
enzymatic function).
Literature Cited
Beach, D., Piper, M., and Nurse, P. 1982. Construction of a Schizosaccharomyces pombe gene bank
in a yeast bacterial shuttle vector and its use to
isolate genes by complementation. Mol. Gen.
Genet. 187:326-329.
Kuo, C.L. and Campbell, J.L. 1983. Cloning of
Saccharomyces cerevisiae DNA replication
genes: Isolation of the CDC8 gene and two genes
that compensate for the cdc8-1 mutation. Mol.
Cell. Biol. 3:1730-1737.
Meeks-Wagner, D., Wood, J.S., Garvik, B., and
Hartwell, L.H. 1986. Isolation of two genes that
affect mitotic chromosome transmission in S.
cerevisiae. Cell 44:53-63.
Rose, M.D. 1987. Isolation of genes by complementation in yeast. Meth. Enzymol. 152:481-504.
Rose, M.D., Novick, P., Thomas, J.H., Botstein, D.,
and Fink, G.R. 1987. A Saccharomyces cerevisiae genomic plasmid bank based on a centromere-containing shuttle vector. Gene 60:237243.
Rothstein, R. 1985. Cloning in yeast. In DNA Cloning, Vol. 2: A Practical Approach (D.M. Glover,
ed.) 4th ed., pp. 45-67. IRL Press, Oxford.
Contributed by Victoria Lundblad
University of California, Berkeley
Berkeley, California
Cloning Yeast
Genes by
Complementation
13.8.4
Supplement 7
Current Protocols in Molecular Biology
Manipulation of Plasmids from Yeast Cells
UNIT 13.9
This unit describes several procedures for manipulating plasmids in yeast cells. The first
(see Basic Protocol 1) is a general method to segregate autonomously replicating plasmids
from cells: plasmid-containing yeast cells are grown in nonselective medium, and
colonies lacking the plasmid are identified by replica plating. The second, plasmid
shuffling (see Basic Protocol 2), represents a specialized version of plasmid segregation
that is useful for analyzing the function of essential genes and for identifying conditional
lethal mutations in essential genes. This method involves introduction of a plasmid
containing mutated versions of essential genes into a strain carrying the wild-type gene
on a URA3 plasmid followed by a genetic selection to remove the URA3 plasmid. The
genetic selection for plasmid loss permits the rapid analysis of many thousands of
colonies. The third approach, plasmid gap repair (see Basic Protocol 3), is based on the
efficient homologous recombination characteristics of yeast cells. Plasmid gap repair can
be be used as a method to incorporate mutagenized DNA fragments into a yeast plasmid,
rescue genomic mutations onto plasmids, or map alleles of a given gene.
SEGREGATION OF PLASMIDS FROM YEAST CELLS
Most E. coli yeast shuttle vectors are lost at a frequency of ∼1% when grown nonselectively. In this simple protocol, a plasmid-bearing yeast strain is grown nonselectively (to
allow segregation of the plasmid) and plated for single colonies. Replica plating onto
selective plates then permits identification of colonies that no longer contain the plasmid.
BASIC
PROTOCOL 1
Materials
Plasmid-bearing yeast strain
YPD or other nonselective liquid medium and plates (UNIT 13.1)
CM dropout plates (UNIT 13.1)
Sterile velvets and replica block (UNITS 13.2 & 1.3)
Additional reagents and equipment for growth and manipulation of yeast (UNIT 13.2)
1. Inoculate several single colonies of a plasmid-bearing yeast strain into individual
10-ml aliquots of nonselective medium (UNIT 13.2) and grow overnight at 30°C.
If no other selection is necessary, YPD medium can be used. If, however, other unstable
genetic markers or plasmids are to be retained, use defined medium supplemented with the
nutrient corresponding to the selectable marker to be lost (e.g., when segregating a LEU2
plasmid, grow in medium supplemented with leucine).
2. Plate for single colonies (UNIT
incubate 2 days at 30°C.
13.2)
on the corresponding nonselective plates and
In most cases, ∼200 to 300 single colonies should be examined (∼100/plate).
3. Replica plate onto selective plates that will identify colonies which have lost the
plasmid-borne selectable marker. Incubate both the selective plates and the nonselective master plate overnight at 30°C.
Those colonies that are present on the master plate but fail to grow on the selective plate
have lost the plasmid.
Saccharomyces
cerevisiae
Contributed by Victoria Lundblad and Heng Zhou
Current Protocols in Molecular Biology (1997) 13.9.1-13.9.6
Copyright © 1997 by John Wiley & Sons, Inc.
13.9.1
Supplement 39
BASIC
PROTOCOL 2
PLASMID SHUFFLING
This technique (based on Boeke et al., 1987) provides a method of analyzing the function
of essential genes as well as a rapid means for identifying conditional lethal mutations in
an essential cloned gene carried on a plasmid (Fig. 13.9.1). In this system, the chromosomal copy of the essential gene is deleted or or otherwise disrupted. Viability is
maintained by the presence of a YEp plasmid carrying an intact copy of this essential
gene, as well as the URA3 gene. Introduction of a second plasmid carrying a temperaturesensitive copy of this gene can relieve selective pressure on the YEp plasmid at the
permissive temperature, generating Ura− derivatives due to loss of the YEp plasmid.
However, at the nonpermissive temperature, the YEp plasmid cannot be lost and no Ura−
segregants are generated. Loss of this plasmid is assayed by replica plating single colonies
onto 5-FOA plates grown at the permissive and nonpermissive temperatures. On these
plates, Ura− segregants appear as 5-FOA-resistant papillae. This replica plating technique
allows a large number of mutagenized clones to be screened rapidly.
Materials
YCp vector (UNIT 13.4) bearing a selectable marker other than URA3
Dropout plates +Ura (UNIT 13.1)
Plates with medium selective for YCp vector and containing 5-FOA (UNIT 13.1)
YPD plates (UNIT 13.1)
YIp5 vector (UNIT 13.4)
Additional reagents and equipment for cloning yeast genes by complementation
(UNIT 13.8), subcloning (UNIT 3.16), mutagenesis (see Chapter 8), lithium acetate
transformation (UNIT 13.7), replica plating (UNIT 13.2), yeast plasmid DNA
preparation (UNIT 13.11), E. coli transformation (UNIT 1.8), and gene replacement
by transplacement (UNIT 13.10)
1. Construct a haploid strain that contains a chromosomal disruption of the essential
gene and a YEp or YCp plasmid carrying both the intact essential gene and URA3 as
a selectable marker (see UNIT 13.8 for a discussion of how to construct such a strain).
A3
UR
YEp
YCp
A B C DE
AB C D E
ts
A E
nonselective
growth
for Ura+
replica plate
onto 5-FOA
YCp
ts
A BCD E
ts
selectable marker
cloned yeast gene
cloned yeast gene with
conditional lethal mutation
disrupted chromosomal
yeast gene
bacterial-derived plasmid DNA
yeast chromosomal DNA
strain containing
ts mutation
A E
23° C
viable
Manipulation of
Plasmids from
Yeast Cells
36 °C
inviable
Figure 13.9.1 Plasmid shuffling is a rapid means of identifying conditional lethal mutations in an
essential cloned gene carried on a plasmid (see accompanying text).
13.9.2
Supplement 39
Current Protocols in Molecular Biology
2. For mutagenesis, subclone (UNIT 3.16) the essential gene onto a YCp vector (bearing a
selectable marker other than URA3) and mutagenize ∼10 to 20 µg using hydroxylamine or other techniques (see Chapter 8). Alternatively, clone mutated derivatives
of an essential gene into the YCp vector.
3. Transform into the strain from step 1 using lithium acetate (UNIT 13.7), selecting for
the YCp plasmid on dropout plates that are supplemented with uracil. Incubate plates
at the permissive temperature (usually 25°C).
This period of growth in the presence of uracil relieves selection for the YEp plasmid, such
that in each transformant colony, a portion of the cells will have lost the plasmid (which
carries the unmutagenized copy of the essential gene). However, note that any transformants carrying a null mutation in this gene on the YCp vector will be unable to lose the
YEp vector.
4. Replica plate (UNIT 13.2) each transformation plate onto two plates that contain 5-FOA
and that still maintain selection for the YCp vector. Incubate one plate at the
permissive temperature and one plate at the nonpermissive temperature (usually
36°C). As a control, replica plate each transformation plate onto two YPD plates and
incubate at the same two temperatures.
Replica plating distinguishes between different categories of transformants by determining
whether a portion of the colony has lost the Ura+ YEp plasmid: such Ura− cells appear as
5-FOAr papillae growing out of the background of the mostly Ura+ replica. Transformants
that carry an unmutagenized copy of the essential gene on the YCp vector will give Ura−
papillae at both temperatures, whereas transformants that contain a temperature-sensitive
mutation in the YCp-borne essential gene will only produce Ura− papillae at the permissive
temperature. Those transformants containing a null mutation in this gene will not generate
papillae at either temperature.
Yeast transformation is often somewhat mutagenic, introducing unlinked chromosomal
mutations in the recipient. Discard all candidates that display a temperature-sensitive
phenotype in the YPD control.
5. Recover from the transformation plate those colonies that gave Ura− papillae at the
permissive temperature but not at the nonpermissive temperature (and which grew
normally at both temperatures on the YPD plates) and streak for single colonies.
6. Retest about six single colonies from each candidate for Ura− papillation at the two
temperatures.
7. For each candidate that retests as a temperature-sensitive mutation in step 6, recover
Ura− papillae from the permissive temperature 5-FOA plate (by streaking out papillae
on a separate plate that maintains selection for the YCp vector). Recover plasmid
from these strains (which now contain only the YCp vector) by isolation of the
plasmid (UNIT 13.11) followed by transformation into E. coli (UNIT 1.8).
8. Subclone (UNIT 3.16) the gene (now carrying a temperature-sensitive mutation) onto a
YIp5 vector, and introduce this mutagenized gene into its chromosomal site using
the transplacement technique described in UNIT 13.10.
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BASIC
PROTOCOL 3
PLASMID GAP REPAIR FOR LOCALIZED MUTAGENESIS AND
ALLELE REPAIR
This gap repair technique protocol is useful for several purposes. First, it can be used as
a method of localized mutagenesis in which a mutagenized DNA fragment (typically
obtained by PCR; UNIT 15.1) can be incorporated into a desired DNA molecule in yeast
cells (Muhlrad et al., 1992). Second, it can be used to introduce a mutation at a specific
position using a mutagenic PCR primer (Brenner et al., 1994). Third, it can be used to
rescue genomic mutations onto a plasmid-borne copy of the same gene for subsequent
analysis. Fourth, it can be used to map multiple alleles of a given gene. In all of these
cases, a gene conversion event initiated by the gapped molecule rescues the introduced
DNA fragment or the chromosomal locus covered by the gapped region; this results in
yeast strains with autonomously replicating plasmids containing the mutagenized DNA
fragment or chromosomal locus of interest (Fig. 13.9.2). The resulting yeast strains can
be directly analyzed for a mutant phenotype, and the plasmids can be rescued and
analyzed.
Materials
YRp or YCp plasmid (UNIT 13.4) with selectable marker
Appropriate restriction enzyme(s) (UNIT 3.1)
Yeast strain with mutation corresponding to selectable marker
Plates with medium selective for the plasmid marker
Additional reagents and equipment for subcloning (UNIT 3.16), PCR (UNIT 15.1), yeast
transformation (UNIT 13.7), growth and manipulation of yeast (UNIT 13.2), and
plasmid segregation (UNIT 13.9)
1. Subclone (UNIT 3.16) the gene of interest into a YRp or YCp plasmid with an appropriate
selectable marker.
2. Gap the plasmid by cutting at two restriction enzyme sites. Gel purify the resulting
plasmid fragment.
An overlap of ∼200 bp on either side of the gap is often used, although as few as 40 bp will
be sufficient.
For introducing a mutation at a specific position, the plasmid may also simply be linearized
rather than gapped; however, contamination with uncut plasmid must be avoided. The gap
or cut must be as close as possible to the mutated site to ensure a high frequency of recovery
of the mutation.
selectable marker
YCp
A B
gapped YCp plasmid DNA
E
PCR insert or chromosomal
gene with mutation ( • )
A BCD E
YCp
A BCD E
Manipulation of
Plasmids from
Yeast Cells
Figure 13.9.2 Plasmid gap repair rescues genomic DNA mutations onto a plasmid-borne copy
of the gene (see accompanying text).
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3. For mutagenesis applications, prepare the desired mutagenized fragment by PCR
(UNIT 15.1). For recovering or mapping chromosomal alleles, proceed to step 4.
A mutagenic PCR reaction using Taq DNA polymerase will introduce mutations at random
positions; mutations at a specific position can be made using a mutagenic primer.
4. Co-transform (UNIT 13.7) 1 µg of gapped plasmid and the PCR-generated fragment
into a yeast strain with appropriate markers. If using gap repair to rescue the genomic
allele, transform with gapped plasmid alone.
Although it is not essential, a 5-fold molar excess of insert is often used.
To ensure that recombination occurs only between the gapped plasmid and the PCR insert,
it is preferable to use a yeast strain deleted for the genomic locus. However, the method
will work even if the genomic locus is present, although a background of undesired
recombination events will arise.
5. Identify successful repair events by plating transformants onto medium selective for
the plasmid marker (UNIT 13.2). For recovering or mapping chromosomal alleles,
identify transformants that are unstable for the selectable marker using the plasmid
segregation technique (UNIT 13.9).
When a mutation is located within the gap, it will be carried on the repaired plasmid.
However if it is next to the gap, the outcome will depend on where the cross-over occurs.
The farther the mutation is from the gapped region, the lower will be the proportion of
plasmids containing the mutation.
6. If using this as a scheme for PCR-based mutagenesis, screen these transformants for
the desired mutant phenotype.
The plasmid can be isolated from yeast (UNIT 13.11) and transformed into E. coli (UNIT 1.8)
for subsequent analysis.
COMMENTARY
Background Information
Techniques for segregating plasmids from
yeast are useful in several experimental situations. For example, after a plasmid containing
a DNA sequence that complements a yeast
mutation has been isolated, it is necessary to
determine whether complementation is due to
sequences present on the plasmid (UNIT 13.8).
This is achieved by demonstrating that the complementation phenotype is lost following segregation of the plasmid.
Plasmid shuffling is a specialized version of
plasmid segregation which is particularly useful for the analysis or mutagenesis of genes that
are essential for cell growth and viability. The
manipulation of such essential genes is complicated by the obvious fact that cells containing
mutated versions of such genes cannot be
propagated. Plasmid shuffling circumvents this
problem by generating a yeast strain in which
the sole copy of an essential gene is present on
a URA3-marked plasmid—i.e., the chromosomal copy of the essential gene is deleted. This
strain is transformed by a second plasmid
(which carries a different marker) containing a
mutated version of the essential yeast gene
(carried on a plasmid with a different marker),
whereupon the URA3 plasmid carrying the
wild-type genes is eliminated (shuffled out) by
replica plating the cells on 5-FOA. This procedure efficiently generates yeast cells that contain only the plasmid with the mutated copy of
the essential gene. If the mutated gene is sufficiently functional to support cell growth, the
colonies can be examined under a variety of
conditions (e.g., low or high temperature). Alternatively, if the mutated gene is nonfunctional, colonies will not arise on the plates
containing 5-FOA.
Although yeast plasmids replicate autonomously from chromosomal DNA, the plasmid
and chromosomal DNAs are similar in virtually
all respects. For this reason, efficient homologous recombination occurs between plasmids
and chromosomes, thereby permitting a number of useful genetic manipulations. In particular, the ends of linearized plasmids can recombine efficiently with introduced DNA fragments or with chromosomal DNA in a process
called plasmid gap repair. This process permits
Saccharomyces
cerevisiae
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Current Protocols in Molecular Biology
Supplement 39
one to incorporate DNA fragments (either wildtype or mutagenized) or genomic alleles into
autonomously replicating plasmids without using standard cloning methods. It is particularly
useful for localized mutagenesis, rescuing
genomic mutations, and genetic mapping (see
Basic Protocol 2).
Literature Cited
Boeke, J.D., Trueheart, J., Natsoulis, G., and Fink,
G.R. 1987. 5-fluoroorotic acid as a selective
agent in yeast molecular genetics. Methods Enzymol. 154:164-175.
Brenner, C., Bevan, A., and Fuller, R.S. 1994. Onestep site-directed mutagenesis. Methods Enzymol. 244:163-165.
Muhlrad, D., Hunter, R.,and Parker, R. 1992. A rapid
method for localized mutagenesis of yeast genes.
Yeast 8:79-92.
Contributed by Victoria Lundblad
Baylor College of Medicine
Houston, Texas
Heng Zhou
Harvard Medical School
Boston, Massachusetts
Manipulation of
Plasmids from
Yeast Cells
13.9.6
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Manipulation of Cloned Yeast DNA
UNIT 13.10
A major advantage of working with yeast is the ability to replace the wild-type chromosomal copy of a gene with a mutant derivative that is constructed in vitro using a cloned
copy of the gene. This technique—unavailable in most other eukaryotes—allows the
phenotype of the mutation to be studied under accurate in vivo conditions, with the
mutation present in single copy at its normal chromosomal location.
Once a cloned gene has been obtained (see UNIT 13.8), there are a variety of techniques
available for introducing a mutant derivative into its corresponding chromosomal site. All
of these methods rely on homologous recombination between the transforming DNA and
the yeast genomic sequences. If the gene has been cloned by complementation, several
of these techniques can be used to demonstrate that the cloned gene corresponds to the
gene defined by the mutation (UNIT 13.8). Gene replacement techniques also allow analysis
of partial or complete gene deletions, which can be introduced into the genome to
determine the null phenotype. In addition, once a cloned gene is available, previously
identified mutant alleles can be recovered via the technique of plasmid gap repair (UNIT
13.9). Gene replacement techniques also allow the creation of modified genes that are
regulated by a desirable (usually inducible) promoter. A specialized version of these
methods permits a general approach for generating conditional alleles by a copper-inducible double shutoff.
INTEGRATIVE TRANSFORMATION
In the steps presented below (based on the procedure described by Hinnen et al., 1978),
a YIp plasmid (UNIT 13.4) harboring both a selectable marker and a cloned gene of interest
is integrated at the chromosomal location of the cloned gene via homologous recombination. The resulting integrant contains the entire plasmid, bracketed by intact copies of
the gene (see Fig. 13.10.1). In the case of cloned genes for which no mutations have been
identified (or where identified mutations have a phenotype that is difficult to score), the
integrated plasmid can be used as a genetic marker—by virtue of the presence of the
selectable gene—for mapping and other genetic studies. Integrative transformation of a
cloned gene can also be used to demonstrate genetic linkage to the mutation used to clone
the gene, providing evidence that the cloned DNA is the corresponding wild-type gene
(discussed in the commentary in UNIT 13.8). It is important to note that this method
introduces a selectable marker at the genomic site of the cloned gene but does not disrupt
the copy of the duplicated gene. Often, multiple tandem integrations can occur, which can
selectable marker
cloned yeast gene
chromosomal yeast gene
bacterial-derived plasmid DNA
yeast chromosomal DNA
double-strand break
recombination site
AB C DE
AB C DE
AB C D E
BASIC
PROTOCOL 1
A BC D E
Figure 13.10.1 Integrative transformation introduces a selectable marker at the chromosomal site
of a cloned gene, via integration of the entire plasmid (see accompanying text).
Contributed by Victoria Lundblad, Grant Hartzog, and Zarmik Moqtaderi
Current Protocols in Molecular Biology (1997) 13.10.1-13.10.14
Copyright © 1997 by John Wiley & Sons, Inc.
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cerevisiae
13.10.1
Supplement 39
be identified by isolating genomic DNA and performing Southern blot analysis (UNITS
13.11, 2.9 & 2.10). In addition, these integrants are unstable and are lost at a frequency of ∼1%
per generation when grown nonselectively.
Although the frequency of integrative transformation is low, it can be increased by
linearizing the plasmid at a restriction site within the cloned gene that is homologous to
the intended site of integration. Linearization also directs, or targets, the recombination
event to the chromosomal homologue of the cloned gene: without a double-strand break
in the cloned gene, the plasmid will integrate at the chromosomal site of either the
selectable gene or the cloned gene, at about equal frequencies. In Figure 13.10.1, the
dashed line indicates the region in which a double-strand break should be introduced.
Materials
YIp shuttle vector (UNIT 13.4)
Appropriate restriction enzyme (UNIT 3.1)
Purified DNA from gene of interest
Carrier DNA
Selective medium (UNIT 13.1)
Additional reagents and equipment for subcloning (UNIT 3.16), yeast transformation
(UNIT 13.7), yeast genomic DNA isolation (UNIT 13.11), and Southern blotting
(UNITS 2.9 & 2.10) or PCR (UNIT 15.1)
1. Subclone (UNIT 3.16) the gene to be studied into a YIp vector.
2. Linearize the plasmid with a restriction enzyme that cuts within the cloned gene.
As long as no other DNA is being cotransformed, it is not necessary to inactivate the
restriction enzyme prior to transformation. If a unique restriction site is not available,
digestion with one or more enzymes to produce a gap in this region is also acceptable, as
long as sufficient homology is present on either side of the gap (>250 bp).
3. Transform an appropriately marked strain with 1 to 10 µg DNA plus carrier DNA
(UNIT 13.7), selecting for the marker present on the plasmid.
4. Purify several transformants on selective medium and isolate genomic DNA (UNIT
13.11). Confirm by Southern hybridization (UNITS 2.9 & 2.10) or PCR (UNIT 15.1) that the
integration has occurred at the desired genomic site and determine whether multiple
integrations have occurred.
GENE REPLACEMENT TECHNIQUES
The four methods described below provide a means of constructing a mutation in vitro in
a cloned gene and reintroducing this mutation at the correct chromosomal site. This allows
assessment of the genetic consequences of a mutation, and is often used to determine
whether or not a gene is essential (by determining if a complete gene deletion is viable).
Two of these techniques—integrative disruption and one-step gene disruption—generate
either insertion or deletion mutations. The third technique—transplacement—is more
generally applicable: it can be used to introduce insertion or deletion mutations containing
a selectable marker, but it can also be used to introduce nonselectable mutations, such as
conditional lethal mutations in an essential gene.
Manipulation of
Cloned Yeast DNA
In each of these procedures, if the goal is to examine whether or not a null mutation is
viable, a diploid strain should be transformed with the appropriate DNA construct. This
allows a potentially lethal mutation to be complemented by the wild-type allele on the
other chromosome. After transformation, the diploid is examined by Southern blot
analysis to confirm that there is one wild-type and one disrupted copy of the gene.
13.10.2
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Current Protocols in Molecular Biology
selectable marker
cloned yeast gene fragment
chromosomal yeast gene
bacterial-derived plasmid DNA
BC D
A B C DE
AB C D
yeast chromosomal DNA
double-strand break
recombination site
B C DE
Figure 13.10.2 Integrative disruption introduces a selectable marker at the chromosomal site of
a cloned gene and simultaneously generates either a 3′ or 5′ deletion in the two copies of the gene
that bracket the plasmid (see accompanying text).
Subsequent sporulation and tetrad dissection (UNIT 13.2) will reveal whether the disruption
is viable. Disruption of an essential gene should result in a ratio of 2+:2− (2 live spores:2
dead spores) in the tetrad analysis; in addition, if a selectable marker (e.g., URA3) is
associated with the disruption, all viable spores should be auxotrophic for this marker
(e.g., Ura−).
Integrative Disruption
This technique (first described by Shortle et al., 1982) generates a deletion in the
chromosomal copy of a cloned gene. An internal fragment of a cloned gene is introduced
into the chromosome on an integrating plasmid. As shown in Figure 13.10.2, this generates
a gene duplication (which brackets the integrated plasmid), but neither copy of the gene
consists of an intact copy: one copy is missing the 3′ end of the gene and one copy is
missing the 5′ end. The procedure is the same as for integrative transformation, with two
exceptions: (1) the starting YIp plasmid, instead of containing an intact gene, contains a
completely internal fragment of the cloned gene, and (2) the strain to be transformed
should be diploid, as explained above.
BASIC
PROTOCOL 2
Two limitations of this technique are that a knowledge of the 5′ and 3′ boundaries of the
coding region of the gene is required and the size of the subcloned fragment must be at
least 250 bp, to promote efficient recombination. However, the disruption can be constructed in one step, without requiring an insertion or insertion/deletion mutation in the
cloned gene, and a selectable phenotype (from the selectable marker on the integrated
YIp plasmid) is associated with the disruption. Like integrative transformation, the
integrants are unstable when grown nonselectively, and multiple integration events can
occur. In addition, linearization of the plasmid increases the frequency of transformation,
as well as targets the integration to the desired site.
Materials
YIp shuttle vector (UNIT 13.4)
Purified DNA from gene of interest
Appropriate restriction enzyme (UNIT 3.1)
Additional reagents and equipment for subcloning (UNIT 3.16) and integrative
transformation (see Basic Protocol 1)
1. Subclone (UNIT 3.16) an internal fragment of the gene into a YIp vector.
2. Linearize the plasmid within this internal fragment by restriction enzyme digestion.
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cerevisiae
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Current Protocols in Molecular Biology
Supplement 39
B
A
ABCD E
A BCD E
ABCD E
ABC DE
disrupted cloned yeast
gene fragment with
selectable marker
chromosomal yeast gene
bacterial-derived plasmid
DNA
yeast chromosomal DNA
recombination site
A BCD E
ABCD E
Figure 13.10.3 One-step gene disruption recombines a fragment of DNA containing a cloned
gene—disrupted by the insertion of a selectable marker—into the chromosomal site of the same
gene (see accompanying text). (A) No portion of the cloned gene fragment is deleted. (B) Sequences BCD are removed from the cloned gene fragment.
3. Continue with transformation and analysis of transformants as for integrative transformation (see Basic Protocol 1, steps 3 and 4).
BASIC
PROTOCOL 3
One-Step Gene Disruption
Like integrative disruption, this method (from Rothstein, 1983) generates a gene disruption in one step via transformation, using a fragment of DNA containing a cloned gene
that is disrupted by a selectable genetic marker. Homologous recombination between the
free DNA ends, which are highly recombinogenic, and homologous sequences in the yeast
genome results in replacement of the wild-type gene by the disrupted copy (see Fig.
13.10.3). The disrupted gene can contain either a simple insertion (of the selectable
marker) or a deletion/insertion mutation. Introduction of these disruptions into the
genome can be achieved in a single step, resulting in stable, nonreverting mutations.
Materials
Purified DNA from gene of interest
Purified DNA from a selectable gene
Appropriate restriction enzyme (UNIT 3.1)
Appropriate yeast strain (e.g., Table 13.10.2)
Additional reagents and equipment for subcloning (UNIT 3.16), gel purification of
DNA (UNIT 2.6), yeast transformation (UNIT 13.7), yeast plasmid DNA isolation
(UNIT 13.11), Southern blotting (UNITS 2.9 & 2.10) or PCR (UNIT 15.1), and tetrad
analysis (UNIT 13.2; optional)
1. Subclone (UNIT 3.16) a suitable selectable gene into the gene of interest, creating in the
process of subcloning a deletion as well, if desired.
Insertion mutations into the cloned gene can also be generated in vivo in E. coli by
transposon mutagenesis. Several transposons have been designed with selectable markers
in E. coli and yeast (Seifert et al., 1986; Huismann et al., 1987).
2. Using appropriate restriction sites, excise a linear fragment that contains the disrupted
gene from the plasmid constructed in step 1 and gel-purify (UNIT 2.6).
Manipulation of
Cloned Yeast DNA
Small amounts of vector sequences (≤200 bp) can be retained on this fragment without
deleterious effects. Ideally, at least 250 bp of the cloned gene should bracket either side of
the inserted selectable gene, to promote recombination at the chromosomal locus of the
cloned gene, rather than at the site of the selectable marker.
13.10.4
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pRS40X
A
S
Left
marker gene X
Right
AS
B
S
Left
marker gene X
Right AS
target gene
S
AS
C
S
Left
marker gene X
PCR product
Right AS
yeast genomic
DNA
yeast genomic
DNA
Figure 13.10.4 PCR mediated one-step gene disruption. (A) The one-step gene-disruption cassette is generated by PCR amplification of marker gene X using primers whose 3′ ends are derived
from the sequences Left and Right found in all pRS40X series vectors. The 5′ 40 nucleotides of
these primers are derived from sequences S (sense strand) and AS (anti-sense strand), which flank
the open reading frame of the target gene. (B) When transformed into yeast, the gene-disruption
cassette can replace the target gene by recombination between the homologous sequences S and
AS. (C) Recombinants can be identified by selection for expression of gene X and confirmed by
subsequent PCR or Southern blot analysis of the target gene locus in the transformants.
3. Transform yeast with 1 to 10 µg of the gel-purified fragment (UNIT 13.7), selecting for
the inserted marker.
4. Confirm the structure of the disruption isolation of genomic DNA (UNIT 13.11) followed
by Southern hybridization (UNITS 2.9 & 2.10) or PCR (UNIT 15.1). If a diploid was
transformed, sporulate and dissect (UNIT 13.2) to obtain haploid spore products with
the disruption (or to observe inviability).
PCR-Mediated One-Step Gene Disruption
One-step gene disruption (first described by Baudin, 1993) is a simple and powerful
technique for removal of specific DNA sequences from the yeast genome. A gene
disruption cassette is constructed that consists of a selectable marker flanked by sequences
derived from the 5′ and 3′ ends of the target gene to be detected. When this linear DNA
fragment is transformed into yeast, homologous recombination between the identical
sequences flanking the marker and target genes results in replacement of the target gene
by the marker gene. In the past, this method suffered from the drawback that a plasmid
carrying the gene disruption cassette had to be constructed before it could be carried out.
Often, suitable restriction sites near the 5′ and 3′ ends of the ORF of the target gene were
not available, making complete deletion of the ORF difficult or impractical. This draw-
ALTERNATE
PROTOCOL 1
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cerevisiae
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Supplement 39
Table 13.10.1 Yeast Markers and Plasmids for
PCR-Mediated One-Step Gene Disruptiona
Gene
Null allele
Template plasmid
URA3
HIS3
LYS2
LEU2
TRP1
MET15
ura3∆0
his3∆200
lys2∆0
leu2∆0
trp1∆63
met15∆0
pRS406
pRS403
pRS317
pRS405
pRS404
pRS401
aAuxotrophic markers and plasmids above are described in Brach-
man et al. (1997) and Sikorski and Hieter (1989).
Table 13.10.2
Suggested Yeast Strains for Gene Replacementa
Strain
ATCC number
Genotype
BY4700
200866
MATa ura3∆
BY4736
BY4727
200898
200889
MATa ade2∆0::hisG his3∆200 met15∆0 trp1∆63 ura3∆0
MATα his3∆200 leu2∆0 lys2∆0 met15∆0 trp1∆63 ura3∆0
aMany other suitable strains are available from the ATCC (strains 200866 to 200902). A recipient diploid can be made
by mating BY4736 and BY4727 and selecting for Ade+ Leu+ Lys+ colonies.
back can be overcome by using PCR to create the gene disruption cassette (see Fig.
13.10.4). Because only very small regions of homology between DNA sequences are
required to direct homologous recombination in yeast, PCR primers that will amplify the
desired auxotrophic marker flanked by 30 to 40 nucleotides of homology to the target
gene can be used to create the gene disruption cassette. The efficiency of this technique
depends upon two factors. First, the homology of the PCR product to the target gene
should be as long as possible, typically 40 nucleotides. Second, this technique works best
if the sequences encoding the auxotrophic marker gene have been completely deleted
from the yeast genome, thus minimizing the chances for homologous recombination
elsewhere in the genome. A set of convenient auxotrophic markers and plasmids for this
technique are shown in Table 13.10.1. Yeast strains carrying different combinations of
these markers (see Brachman et al., 1997) are available from the ATCC (strains 200866
to 200902); some suggested strains are described in Table 13.10.2. Using the pRS40X
series of vectors as the template for PCR, one set of PCR primers is sufficient to make a
gene knockout marked by any one of the six different auxotrophic markers listed in Table
13.10.1.
Manipulation of
Cloned Yeast DNA
Materials
10× PCR buffer (e.g., as supplied with Taq DNA polymerase from Boehringer
Mannheim)
100 mM Tris⋅Cl/15 mM MgCl2/500 mM KCl, pH 8.3 (20°C)
4dNTP mixture (UNIT 3.4)
pRS40X-series vector (e.g., Table 13.10.1)
pRS Left and Right primers (see step 1)
Taq DNA polymerase
Appropriate yeast strain (e.g., Table 13.10.2)
Selective medium (UNIT 13.1)
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Additional reagents and equipment for PCR (UNIT 15.1), yeast transformation
(UNIT 13.7), yeast genomic DNA isolation (UNIT 13.11) and Southern blotting
(UNITS 2.9 & 2.10; optional)
1. Design PCR primers for the disruption cassette as follows (see Fig. 13.10.4). Pick 40
nucleotides of sequence from the sense strand at the 5′ end of the target gene ORF
and add to the 5′ end of one of the two sequences given below. Similarly, pick 40
nucleotides of sequence from the anti-sense strand at the 3′ end of the target-gene
ORF to be deleted and add to the 5′ end of the other sequence listed below (found in
all pRS-series vectors). Obtain primer from oligonucleotide supplier.
Left primer:
Right primer:
5′-CTGTGCGGTATTTCACACCG-3′
5′-AGATTGTACTGAGAGTGCAC-3′
This results in two 60-nucleotide primers sufficient to amplify any of the auxotrophic
markers in the pRS-series vectors. The overall primer length is set at 60 nucleotides because
this is the longest primer length that is convenient and economical to obtain commercially,
and because primers of this length typically give satisfactory results.
2. Set up several 100-µl PCR reactions containing:
10 µl 10× PCR buffer
200 µM 4dNTP mixture
100 pmol of each Left and Right primer
10 ng pRS40X
2.5 U Taq DNA polymerase.
pRS40X series vectors are used as the template because they lack sequences that allow
their stable maintenance in yeast, thus obviating need for purification of the PCR product
away from the template DNA. Because the number of transformants obtained from a yeast
transformation is proportional to the amount of input DNA, it is worthwhile to perform
several 100-ìl reactions, combine and concentrate them by ethanol precipitation (UNIT 2.1),
and then transform all of the DNA together.
3. Perform PCR amplification (UNIT 15.1) using the following cycling conditions:
1 cycle:
30 cycles:
1 cycle:
2 min
30 sec
1 min
2 min
10 min
94°C
94°C
55°C
72°C
72°C
(denaturation)
(denaturation)
(annealing)
(extension)
(extension).
4. Transform (UNIT 13.7) the PCR product into an appropriate yeast strain and select for
expression of the auxotrophic marker.
Because the transformation efficiency of these PCR products is low, it is critical that a
high-efficiency transformation procedure—such as lithium acetate transformation with
early log-phase cell (see UNIT 13.7)—be used. Also, if the gene being deleted is essential for
life, the deletion mutation must be created in a diploid strain.
5. Isolate genomic DNA (UNIT 13.11) and identify strains carrying the gene disruption by
PCR (UNIT 15.1) with primers that flank the expected mutation, or by Southern blot
analysis (UNITS 2.9 & 2.10).
As noted above, the efficiency of this procedure is dependent upon the complete absence
of homology to the auxotrophic marker in the recipient strain. In cases where substantial
homology to the auxotrophic marker remains, the frequency of transformants with the
expected deletion may drop to as low as a few percent. In such cases, the deletions may
still be found by screening many clones by PCR or, if possible, by first identifying potential
deletion strains among the transformants by some expected mutant phenotype.
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cerevisiae
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Current Protocols in Molecular Biology
Supplement 39
UR
cloned yeast gene
with mutation (•)
A3
chromosomal yeast gene
AB C DE
bacterial-derived plasmid DNA
yeast chromosomal DNA
AB C DE
double-strand break
recombination site
AB C D E
URA 3
ABC D E
select for 5-FOAr
AB C D E
or
ABC D E
Figure 13.10.5 Transplacement allows the introduction of any mutation in a cloned gene that does
not have a selectable phenotype (see accompanying text).
BASIC
PROTOCOL 4
Transplacement
This method (based on Scherer and Davis, 1979), also called allele replacement, provides
a general means of introducing any type of mutation constructed in vitro that does not
have a selectable phenotype into its corresponding chromosomal location. It can be used
either to introduce a single defined mutation or to screen a mutagenized collection of
plasmids. The mutated gene is introduced into yeast on a YIp plasmid, which usually
contains the URA3 gene as a selectable marker. After transformation, selection for Ura+
results in an integration event, such that mutant and wild-type alleles bracket the
URA3-containing plasmid sequences (Fig. 13.10.5). Subsequent eviction of the plasmid
(containing one copy of the gene) is monitored by screening for colonies that are URA−
and therefore resistant to the drug 5-fluoroorotic acid (5-FOA). The 5-FOAr colonies are
then screened for the desired mutant phenotype; the proportion of 5-FOAr colonies
containing the mutant allele will depend on the position of the mutation relative to the
length of the flanking homologous DNA.
Materials
YIp shuttle vector (UNIT 13.4)
Purified DNA from gene of interest
Nonselective liquid medium (YPD or uracil-containing minimal medium; UNIT 13.1)
5-FOA plates (UNIT 13.1)
Additional reagents and equipment for subcloning (UNIT 3.16), integrative
transformation (see Basic Protocol 1), yeast genomic DNA isolation (UNIT 13.11),
and Southern blotting (UNITS 2.9 & 2.10) or PCR (UNITS 15.1.)
1. Subclone (UNIT 3.16) the gene of interest into YIp5.
2. Mutagenize the plasmid as desired, either via the techniques presented in Chapter 8,
or by introducing defined deletion or insertion mutations.
3. Proceed with linearization, transformation, and analysis of transformants as for
integrative transformation (see Basic Protocol 1, steps 2 to 4).
Manipulation of
Cloned Yeast DNA
4. Grow transformants overnight in liquid medium without selection. Plate for single
colonies on nonselective medium (∼100 single colonies) and grow 2 days at 30°C.
13.10.8
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Current Protocols in Molecular Biology
marker
integrating plasmid
promoter
chromosomal locus
TAG 5′ ORF
ORF
result of integration
marker
promoter TAG
5′ ORF
entire ORF
Figure 13.10.6 A single integrative transformation step introduces the desired allele at the normal
genome locus and truncates the wild-type allele.
Alternatively, cells can be plated or streaked directly on 5-FOA plates, in which case
proceed to step 6.
If the goal is to identify conditional lethal mutants, a large number of transformants should
be examined (since some proportion of transformants will have received unmutagenized
plasmid and since conditional lethal mutations can be rare events). In addition, this
nonselective period of growth should be at the permissive temperature (usually 23°C).
5. Replica plate onto 5-FOA plates and grow overnight to identify Ura− segregants.
Colonies that have evicted the plasmid during the period of growth in liquid medium will
give completely resistant replicas on the 5-FOA plates; these colonies should be recovered
from the nonselective plate (rather than from the 5-FOA plate) and used in step 6, below.
Other colonies that have evicted the plasmid during the period of growth of the colony will
appear “patchy” on the 5-FOA replica (often referred to as papillation).
6. Screen 5-FOAr colonies for the presence of a mutant phenotype. If the mutation is
expected to alter the restriction pattern, confirm the genotype of mutant colonies in
comparison to wild-type by isolation of genomic DNA (UNIT 13.11) followed by
Southern hybridization (UNITS 2.9 & 2.10) or PCR (UNIT 15.1).
CREATING MODIFIED GENES BY ONE-STEP INTEGRATIVE
REPLACEMENT
BASIC
PROTOCOL 5
This method may be used to create a yeast strain in which a gene of interest is controlled
by any desired (usually inducible) promoter. The same method may be used to create an
N-terminal fusion to any chosen epitope tag or other cassette. A single integration event
introduces the new allele of the gene at its normal locus and simultaneously truncates the
normal endogenous copy of the gene, rendering it nonfunctional (Fig. 13.10.6).
Materials
Purified DNA from gene of interest
Yeast integrating shuttle vector with selectable marker
Appropriate restriction enzyme (UNIT 3.1)
Appropriate yeast strain
Selective medium (UNIT 13.1)
Saccharomyces
cerevisiae
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Current Protocols in Molecular Biology
Supplement 39
Additional reagents and equipment for subcloning (UNIT 3.16), yeast transformation
(UNIT 13.7), yeast genomic DNA isolation (UNIT 13.11), and Southern blotting
(UNITS 2.9 & 2.10) or PCR (UNIT 15.1)
1. Select a short 5′ fragment of the open reading frame (ORF) of the gene of interest.
This fragment should:
a. begin at the first codon of the gene;
b. be too short to encode a functional product; in addition, its short product should
not have the potential to interfere with the normal activity of the full-length gene
product;
c. be long enough (preferably ≥300 bp) to be efficiently targeted by homology to
the corresponding chromosomal region; and
d. contain a restriction site (to be used for targeting the final integrating plasmid to
the correct locus) that will remain unique in the final construct and that leaves
reasonable stretches of homology to the target gene on both sides of the double
strand break.
In order to minimize the chances of undesired integration at the promoter locus, it is
preferable for the restriction site to be located slightly closer to the 3′ end of the ORF
fragment than to its 5′ end.
2. Subclone (UNIT 3.16) this 5′ fragment downstream of the desired promoter in an
integrating yeast shuttle plasmid. If desired, this construct may also include an epitope
tag (or other fusion cassette) appended in frame to the N terminus of the ORF
fragment.
This subcloning is usually most easily accomplished by PCR. Be certain to situate the initial
ATG in the appropriate promoter context. Occasionally, an N-terminal epitope tag may
interfere with proper function of a gene. In these situations it is advisable to try subcloning
the ORF fragment downstream of the promoter with no N-terminal fusion. Alternatively, a
modification of this procedure (using a short, nonfunctional 3′ piece of the ORF fused
upstream of the desired tag) may be used to create a C-terminal fusion to an epitope tag.
3. Linearize the construct at a unique restriction site within the ORF fragment.
If no restriction site in the ORF fragment is unique, a partial digest may be used to create
a mixed population of linearized molecules for transformation. A correspondingly larger
number of transformants should then be screened in step 6 for correctly targeted integration. On the other hand, if no restriction sites are present in the ORF fragment, it may be
necessary to introduce one by silent PCR mutagenesis (UNIT 8.5).
4. Transform the linearized molecule into an appropriate yeast strain (UNIT 13.7).
5. Plate the transformations onto medium selecting for the plasmid marker (UNIT 13.2).
Remember that if placing an essential gene under the control of an inducible promoter, it
is important to plate the cells onto inducing medium.
6. Isolate genomic DNA (UNIT 13.11) and identify clones containing the modified genes
by Southern blotting (UNITS 2.9 & 2.10), genomic PCR (UNIT 15.1), or phenotypic
assessment.
When working with strains created by this method, be sure to maintain selection for the
plasmid marker in order to prevent looping out of the integrated plasmid.
Manipulation of
Cloned Yeast DNA
13.10.10
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Current Protocols in Molecular Biology
CREATING MODIFIED GENES BY TRANSPLACEMENT
For some genes, it may be difficult or impossible to select a 5′ ORF fragment that is long
enough for efficient integration yet too short to retain function. In these situations,
inducible alleles may be introduced by complete allele replacement (see Basic Protocol
4). Such replacements are stable, leave no short 5′ ORF fragment upstream, and avoid the
need to maintain selection for the plasmid marker. The allele replacement may be
accomplished as follows:
ALTERNATE
PROTOCOL 2
1. Create a molecule as for one-step integrative replacement (see Basic Protocol 5, steps
1 and 2), being sure to use an integrating vector with URA3.
2. Just upstream of the promoter, subclone (UNIT 3.16) a fragment (usually ≥500 bp) of
the 5′ flanking region of the target gene.
When selecting this fragment, take care to preserve the uniqueness of the targeting
restriction site in the ORF fragment.
3. Linearize the molecule and integrate into yeast (see Basic Protocol 5, steps 3 to 5).
4. Streak the URA+ transformants onto 5-FOA plates (see Basic Protocol 4, step 5).
5. Screen the 5-FOAr clones obtained for the correctly replaced allele (see Basic
Protocol 4, step 6).
CREATION OF CONDITIONAL ALLELES BY COPPER-INDUCIBLE
DOUBLE-SHUTOFF PROCEDURE
BASIC
PROTOCOL 6
This method (first described in Moqtaderi et al., 1996), which is useful in the functional
analysis of essential genes, allows the creation of conditional expression strains in which
the addition of copper causes expression of a gene of interest to be shut off at both the
RNA and the protein levels. The RNA shutoff is accomplished by harnessing a transcriptional repressor, and the protein degradation is achieved by exploitation of the N-end rule,
whereby a protein’s rate of turnover is dictated by its N-terminal amino acid (Varshavsky,
1992). Traditional depletion systems generally rely on the removal of positive agents or
on a complete change in medium (e.g., galactose to glucose shift), requiring the cells to
be washed at the time of the shift to nonpermissive conditions. In contrast, this method
works indirectly, by the induction of negative agents (the simple addition of copper to
stimulate the production of a transcriptional repressor and a protein degradation factor),
which obviates the need to wash the cells.
Specifically, the parent strain used must harbor copper-inducible alleles of ROX1, a
transcriptional repressor, and of UBR1, which encodes the N-end recognition component
of the ubiquitin protein degradation pathway. In this background, the gene of interest is
replaced with an allele fused to an N-end recognition sequence for rapid degradation and
driven by the ANB1 promoter, a natural target of Rox1. In the absence of copper, neither
Rox1 nor Ubr1 is produced, and the target gene is expressed. The addition of copper
results in rapid activation of both ROX1 and UBR1, leading to the simultaneous repression
of the target gene’s transcription and rapid degradation of its existing protein product
(Figs. 13.10.7 and 13.10.8).
Materials
Plasmid ZM168, containing the ANB1 promoter driving a
ubiquitin-arginine-lacI-HA (URLF) cassette immediately followed by a
polylinker
Yeast integrating shuttle vector with selectable marker
Yeast strain ZMY60 (MATa, ade2-101, HIS+, LEU+, trp1∆1, ura3-52)
Saccharomyces
cerevisiae
13.10.11
Current Protocols in Molecular Biology
Supplement 39
ANB1
ANB1
target gene
URLF
represses RNA synthesis
ROX1
+
target RNA
add Cu2+
+
ANB1
degrades tagged protein
UBR1
target protein
Figure 13.10.7 A two-pronged conditional knock-out strategy shuts off the target gene’s expression at both the RNA and the protein levels.
marker
integrating plasmid
ANB
URLF
5′ ORF
chromosomal locus
ORF
result of integration
marker
5′ ORF
ANB
URLF
entire ORF
Figure 13.10.8 Introduction of URLF-tagged, ANB1-driven allele of the desired target gene into
the genome.
Centromeric plasmid bearing the same marker as the integrating vector
Synthetic medium plates selecting for the integrated plasmid marker (UNIT 13.1),
with and without 500 µM cupric sulfate (CuSO4)
1. Following the guidelines provided for one-step replacement (see Basic Protocol 5,
steps 1 and 2), select a short, nonfunctional 5′ fragment of the intended target gene,
and subclone it (UNIT 3.16) into the polylinker of the plasmid ZM168 in frame with the
URLF cassette.
2. Subclone the entire ANB1-URLF-ORF fragment into a yeast integrating shuttle
vector bearing the preferred selectable marker.
Take care to maintain the uniqueness of the targeting restriction site in the ORF.
3. Linearize the final construct at the unique restriction site within the ORF fragment
and transform it (UNIT 13.7) into yeast strain ZMY60. In parallel, transform ZMY60
Manipulation of
Cloned Yeast DNA
13.10.12
Supplement 39
Current Protocols in Molecular Biology
with a centromeric plasmid bearing the same yeast marker as the final integrating
construct. Plate onto synthetic medium selecting for the plasmid marker.
A different strain background may be desired. If this is the case, it is possible to generate
another strain containing stably integrated, copper-inducible alleles of ROX1 and UBR1
by using the two-step gene replacement constructs ZM195 and ZM197. (See Basic Protocol
4 for details of how to construct stable gene replacements.)
4. Streak the transformants onto (a) synthetic medium selecting for the integrated
plasmid marker with no added copper and (b) synthetic medium selecting for the
integrated plasmid marker and containing 500 µM CuSO4. Grow plates 2 days at
30°C.
When preparing solid medium for this purpose, it is generally advisable to add all necessary
amino acids and the copper when pouring the plates. Attempts to spread liquid amino acid
supplements or copper on top of a previously poured plate may result in an uneven copper
concentration, making strain growth patterns more difficult to interpret.
5. Analyze transformant growth to assess integration. If the target gene is essential,
correct integrants should fail to grow on copper-containing medium. Transformants
carrying the centromeric plasmid should grow normally on both media.
If the depletion is inefficient, the copper concentration may be raised to 1 mM, but beyond
this level cell growth may begin to be impaired due to copper toxicity. If the target gene is
not essential, test for an expected phenotype in the presence of copper or analyze the
transformants by isolating genomic DNA (UNIT 13.11) and Southern blotting (UNITS 2.9 &
2.10).
6. For time-course experiments, grow both the conditional strain and the centromeric
plasmid control strain in liquid synthetic medium without copper but selective for
the plasmid marker. Add 500 µM CuSO4 to initiate depletion and harvest cells at
varying time-points for analysis.
A standard procedure is to assay the cellular process under investigation at time-points
over the next several hours after the addition of copper (usually up to 8 hr) to test the effects
of increasing levels of depletion. The centromeric plasmid control strain is included to
represent a strain expressing a wild-type level of the gene product of interest. The
conditional strain, even under permissive conditions, harbors an allele of the target gene
driven by a heterologous promoter, and it therefore may not yield a truly wild-type level of
expression. It is important to maintain selection for the marker at all times to avoid loss of
the integrated plasmid. Even after the addition of copper, it is normal for the cells to
continue to grow for several hours. The need to maintain the cells in exponential growth
throughout most experiments thus makes it inadvisable to add copper at too high a cell
density; a typical OD600 at the time of shift is ∼0.25.
LITERATURE CITED
Baudin, A., Ozier-Kalogeropoulos, O., Denouel, A.,
Lacroute, F., and Cullin, C. 1993. A simple and
efficient method for direct gene deletion in Saccharomyces cerevisiae. Nucl. Acids Res.
21:3329-3330.
Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E.,
Li, J., Hieter, P., and Boeke, J. 1998. Designer
deletion strains derived from Saccharomyces
cerevisiae S288C: A useful set of strains and
plasmids for PCR-mediated gene disruption and
other applications. Yeast 14:115-132.
Hinnen, A., Hicks, J.B., and Fink, G.R. 1978. Transformation of yeast. Proc. Natl. Acad. Sci. U.S.A..
75:1929-1933.
Huismann, O., Raymond, W., Foelich, K., Errada,
P., Kleckner, N., Botstein, D., and Hoyt, M.A.
1987. A Tn10-lacZ-kanR-URA3 gene fusion
transposon for insertion mutagenesis and fusion
analysis of yeast and bacterial genes. Genetics
1116:191-199.
Moqtaderi, Z., Bai, Y., Poon, D., Weil, P.A., and
Struhl, K. 1996. TBP-associated factors are not
generally required for transcriptional activation
in yeast. Nature 383:188-191.
Rothstein, R.J. 1983. One-step gene disruption in
yeast. Methods Enzymol. 101:202-210.
Saccharomyces
cerevisiae
13.10.13
Current Protocols in Molecular Biology
Supplement 39
Scherer, S. and Davis, R.W. 1979. Replacement of
chromosome segments with altered DNA sequences constructed in vitro. Proc. Natl. Acad.
Sci. U.S.A. 76:4951-4955.
Seifert, H.S., Chen, E.Y., So, M., and Heffron, F.
1986. Shuttle mutagenesis: A method of
transposon mutagenesis for Saccharomyces
cerevisiae. Proc. Natl. Acad. Sci. U.S.A. 83:735739.
Shortle, D., Haber, J.E., and Botstein, D. 1982.
Lethal disruption of the yeast actin gene by integrative DNA transformation. Science 217:371373.
Sikorski, R.S. and Hieter, P. 1989. A system of
shuttle vectors and yeast host strains designed for
efficient manipulation of DNA is Saccharomyces
Cerevisiae. Genetics 122:19-27.
Contributed by Victoria Lundblad
Baylor College of Medicine
Houston, Texas
Grant Hartzog (PCR-mediated one-step
gene disruption)
Harvard Medical School
Boston, Massachusetts
Zarmik Moqtaderi (creation of modified
genes and conditional alleles)
Harvard Medical School
Boston, Massachusetts
Varshavsky, A. 1992. The N-end rule. Cell 69:725735.
Manipulation of
Cloned Yeast DNA
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Current Protocols in Molecular Biology
PREPARATION OF YEAST DNA, RNA,
AND PROTEINS
SECTION IV
Preparation of Yeast DNA
UNIT 13.11
Molecular studies in yeast often require the isolation of both plasmid and chromosomal
yeast DNA. Plasmid DNA is used in the transformation of E. coli, whereas chromosomal
DNA is used for Southern hybridization analysis, in vitro amplification by the polymerase
chain reaction (PCR), or cloning of integrated plasmids. This unit presents two variations
of the “smash and grab” protocol (Hoffman and Winston, 1987) that produce suitable
DNA for all these applications. These protocols work for both Saccharomyces cerevisiae
and Schizosaccharomyces pombe, although for S. pombe a suitable liquid medium must
be used to grow the cells.
RAPID ISOLATION OF PLASMID DNA FROM YEAST
Plasmid DNA is released from a yeast transformant along with chromosomal DNA in a
rapid, 10-min protocol by vortexing with glass beads in the presence of detergents, phenol,
chloroform, and isoamyl alcohol. After centrifugation to pellet cell debris, no further
purification of the DNA is required for transformation of competent E. coli cells.
BASIC
PROTOCOL
Materials
YPD or appropriate selective medium (UNIT 13.1)
Yeast colony containing the plasmid of interest
Breaking buffer (see recipe)
25:24:1 (v/v/v) phenol/chloroform/isoamyl alcohol (with buffered phenol; UNIT 2.1)
0.45- to 0.52-mm acid-washed glass beads (UNIT 13.12; Thomas Scientific)
Competent E. coli cells HB101 or MH1 (UNIT 1.8)
LB plates (UNIT 1.1) containing appropriate antibiotic (UNIT 1.4)
13 × 100–mm glass tubes, sterile
30°C incubator with shaker or roller drum
Additional reagents and equipment for growth of E. coli on solid medium
(UNIT 1.3) and transformation of E. coli (UNIT 1.8)
1. Inoculate 2 ml medium in a 13 × 100–mm sterile glass tube with a single yeast colony
containing the plasmid of interest. Grow overnight to stationary phase at 30°C in
either a roller drum or a shaking incubator.
Transformants carrying stable plasmids (UNIT 13.4) should be grown in YPD to get a dense
culture. Transformants carrying unstable plasmids should be grown in selective medium
to increase the percentage of plasmid-containing cells.
2. Transfer 1.5 ml of the overnight culture to a microcentrifuge tube and spin 5 sec at
high speed, room temperature. Pour off supernatant and disrupt pellet by vortexing
briefly.
3. Resuspend cells in 200 µl breaking buffer. Add 0.3 g glass beads (∼200 µl volume)
and 200 µl phenol/chloroform/isoamyl alcohol. Vortex 2 min at highest speed.
The amount of vortexing required can vary depending upon the vortex used. Determine by
microscopic examination the minimum vortexing required for a particular machine to break
80% to 90% of the cells.
For simultaneous preparation of DNA from several yeast strains, a multi-tube vortexer or
a multi-tube head for a standard vortexer can be used (VWR Scientific) and vortexing time
should be increased to 3 min.
Contributed by Charles S. Hoffman
Current Protocols in Molecular Biology (1997) 13.11.1-13.11.4
Copyright © 1997 by John Wiley & Sons, Inc.
Saccharomyces
cerevisiae
13.11.1
Supplement 39
4. Microcentrifuge 5 min at high speed, room temperature.
5. Transform competent E. coli HB101 or MH1 with 1 to 2 µl of the aqueous layer. Plate
on LB plates containing the appropriate antibiotic to select for the drug-resistance
marker on the plasmid.
See Critical Parameters and Troubleshooting for guidelines on transformation.
Save 50 ìl of the aqueous layer and store at −20°C in case additional transformations are
required.
ALTERNATE
PROTOCOL
RAPID ISOLATION OF YEAST CHROMOSOMAL DNA
The DNA preparation described in the Basic Protocol can be easily scaled up to prepare
chromosomal DNA for use in Southern hybridization analysis (UNIT 2.9A), in vitro DNA
amplification by PCR (UNIT 15.1), or restriction digestion and ligation (UNITS 3.1 & 3.16) to
clone integrated plasmids. This procedure is significantly faster than other protocols used
to isolate high-molecular-weight DNA. Although the DNA is subject to shearing in this
protocol, the resulting DNA is of sufficiently high molecular weight to enable detection
of 19-kb restriction digestion products by Southern hybridization analysis.
Additional Materials (also see Basic Protocol)
TE buffer (APPENDIX 2)
1 mg/ml DNase-free RNase A (UNIT 5.5)
4 M ammonium acetate solution (APPENDIX 2)
100% ethanol
18 × 150–mm glass culture tubes or 17 × 100–mm disposable
polypropylene tubes, sterile
Tabletop centrifuge
Prepare yeast cells
1. Grow a 10-ml culture of yeast in YPD overnight to stationary phase in either 18 ×
150–mm sterile glass culture tubes or in 17 × 100–mm sterile, polypropylene tubes
(see Basic Protocol, step 1).
2. Spin culture 5 min in a tabletop centrifuge at 1200 × g (3000 rpm), room temperature.
Aspirate or pour off supernatant, and resuspend cells in 0.5 ml water.
3. Transfer the resuspended cells to a microcentrifuge tube and spin 5 sec at room
temperature. Pour off supernatant and disrupt pellet by vortexing briefly.
This wash step in water removes any remaining medium.
Break open the cells
4. Resuspend cells in 200 µl breaking buffer. Add 0.3 g glass beads (∼200 µl volume)
and 200 µl phenol/chloroform/isoamyl alcohol and vortex at highest speed for 3 min.
The amount of vortexing required can vary depending upon the vortex used. Determine by
microscopic examination the minimum vortexing required for a particular machine to break
80% to 90% of the cells.
If using a multi-tube vortex rather than a single-tube vortex, the time of vortexing may need
to be increased up to 4 or 5 min to get more efficient breakage of cells. However, longer
vortexing can result in shearing of DNA molecules.
5. Add 200 µl TE buffer and vortex briefly.
Preparation of
Yeast DNA
6. Microcentrifuge 5 min at high speed, room temperature, and transfer aqueous layer
to a clean microcentrifuge tube. Add 1 ml of 100% ethanol and mix by inversion.
13.11.2
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Current Protocols in Molecular Biology
7. Microcentrifuge 3 min at high speed, room temperature. Remove supernatant and
resuspend pellet in 0.4 ml TE buffer.
Degrade RNA contaminants and recover DNA
8. Add 30 µl of 1 mg/ml DNase-free RNase A, mix, and incubate 5 min at 37°C.
9. Add 10 µl of 4 M ammonium acetate and 1 ml of 100% ethanol. Mix by inversion.
10. Microcentrifuge 3 min at high speed, room temperature. Discard supernatant and dry
pellet. Resuspend DNA in 100 µl TE buffer.
Yields of ∼20 ìg of chromosomal DNA should be obtained. This DNA is ready to use for
restriction digestion (UNIT 3.1), in vitro PCR amplification (UNIT 15.1), or Southern blot
analysis (UNIT 2.9). For Southern blots, best results are obtained when 5 ìl DNA (∼1 ìg) is
digested in a total volume of 20 ìl. To amplify by PCR, 2 ìl of DNA should be used in a
50-ìl reaction.
REAGENTS AND SOLUTIONS
Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see
APPENDIX 2; for suppliers, see APPENDIX 4.
Breaking buffer
2% (v/v) Triton X-100
1% (v/v) sodium dodecyl sulfate (SDS)
100 mM NaCl
10 mM Tris⋅Cl, pH 8.0
1 mM EDTA, pH 8.0
Store ≤1 year at room temperature
COMMENTARY
Background Information
These DNA isolation protocols simultaneously break yeast cells by vortexing with glass
beads in a detergent solution and separate nucleic acids from protein by phenol/chloroform
extraction. They differ in the amount of additional manipulation required to obtain DNA of
the quality needed for a particular procedure.
Other protocols also describe the isolation of
high-molecular-weight DNA (Cryer et al.,
1975; Winston et al., 1983; Davis et al., 1980).
However, even the protocols designed to be
rapid DNA preparations require >3 hr time. In
general, the protocols described here produce
DNA of sufficient molecular weight for any
desired application, except for the production
of insert DNA for the construction of genomic
libraries.
Many situations require the isolation of plasmid DNA from a yeast transformant to transform E. coli. Examples include cloning of (1)
the wild-type copy of a gene by complementation of a recessive mutant (UNIT 13.8), (2) a
dominant mutant allele of a gene by transformation of a wild-type strain, or (3) a mutant
allele of a gene by plasmid gap repair (UNIT 13.10).
Plasmid DNA is also needed to identify a de-
sired mutant allele of a gene by the plasmid
shuffle technique (UNIT 13.10). Most yeast plasmids used in molecular biology are shuttle
vectors carrying a selectable marker and an
origin of replication (ori) for transformation of
and maintenance in E. coli (UNIT 13.4). Because
significantly more plasmid DNA can be isolated from an E. coli transformant than from a
yeast transformant, E. coli is used to amplify
the DNA for restriction mapping and other in
vitro manipulations of the DNA as well as for
retransformation of yeast strains.
Other situations require the isolation of
chromosomal DNA. The most common use of
this DNA is for Southern hybridization analyses. Such analyses are needed to confirm that
integration of a YIp plasmid or of a marked gene
disruption has occurred by homologous recombination. Chromosomal DNA can also be used
as a template for in vitro amplification by PCR
in place of Southern analyses to determine
whether DNA is integrated by homologous
recombination. PCR can also be used in place
of gap repair to clone mutant alleles of a given
gene. Finally, chromosomal DNA is needed to
clone DNA adjacent to an integrated plasmid.
This is done by digesting the chromosomal
Saccharomyces
cerevisiae
13.11.3
Current Protocols in Molecular Biology
Supplement 39
Preparation of
Yeast DNA
DNA with a restriction enzyme that creates a
restriction fragment containing the plasmid origin of replication and selectable marker, as well
as the flanking DNA of interest (Southern
analyses are required to determine the appropriate restriction enzyme for this construction).
Ligation of this yeast DNA, followed by transformation of E. coli, results in the isolation of
the cloned flanking DNA, as only the appropriate piece of DNA will confer drug resistance
upon the E. coli strain.
These protocols do not produce intact chromosomal DNA. Yeast chromosomes can be
isolated by methods involving gentle lysis of
cells in agarose blocks (UNIT 2.5B; Carle and
Olson, 1987). Intact chromosomes can be used
to map a cloned gene to a given chromosome
by Southern hybridization analysis or as a size
standard for field-inversion or orthogonalfield-alteration gel electrophoresis (UNIT 2.5B).
Three minutes is generally sufficient, but this
time can vary from vortex to vortex. Too much
vortexing can result in a weak signal for restriction fragments <10 kb. The other consideration
for optimizing Southern blot analysis is the
amount of DNA used per digest. This protocol
generates a very consistent yield of readily
digested DNA. Increasing the amount of DNA
per digest can result in incomplete cutting and
is not recommended.
Critical Parameters and
Troubleshooting
Time Considerations
There is a significant difference among E.
coli strains regarding their transformation efficiency with plasmid DNA isolated by this protocol. One commonly used strain, HB101 (Table 1.4.5), works well with either CaCl2 transformation or electroporation protocols (UNIT
1.8). A less commonly used strain, MH1 (Hall
et al., 1984), is even better suited for these
transformations when made competent by the
RbCl method (Hanahan, 1983). Because MH1
is a derivative of MC1061, it is possible that
this strain would also transform well using the
Hanahan protocol. Many other strains do not
work as well, even when they appear to be more
competent as judged by transformation with a
control plasmid used to standardize transformation efficiency. Therefore, problems with
transformations may be due to the choice of the
E. coli host strain.
It is important to note that only 1 or 2 µl of
the aqueous layer should be used per transformation, as larger amounts are increasingly toxic
to E. coli. Attempts to purify the DNA with
ethanol precipitations are generally unsuccessful, possibly due to co-precipitation of material
that inhibits transformation. A 1:1 isopropanol
precipitation can be effective in concentrating
the DNA when the original attempt fails to
produce transformants. In general, however,
this additional manipulation is unwarranted.
When preparing DNA for use in Southern
hybridization analysis, it is important to minimize the vortexing time needed to break the
cells in order to reduce shearing of the DNA.
Anticipated Results
The basic protocol for isolating plasmid
DNA from yeast transformants gives sufficient
DNA for transformation of E. coli. The number
of transformants obtained is a function of copy
number of plasmid and transformation efficiency of E. coli host strain. The protocol for
isolating chromosomal DNA yields ∼20 µg
DNA from a 10-ml stationary-phase culture.
It should take ∼10 min to prepare plasmid
DNA for transformation of E. coli. The
genomic DNA preparation takes from 45 min
to 1 hr for twelve cultures. The vortexing step
can become much more time-consuming if only
a single-tube vortex is available, as only three
microcentrifuge tubes can be vortexed simultaneously.
Literature Cited
Carle, G.F. and Olson, M.V. 1987. Orthogonal-fieldalternation gel electrophoresis. Methods Enzymol. 155:468-482.
Cryer, D.R., Eccleshal, R., and Marmur, J. 1975.
Isolation of yeast DNA. Methods Cell Biol.
12:39-44.
Davis, R.W., Thomas, M., Cameron, J., St. John,
T.P., Scherer, S., and Padgett, R.A. 1980. Rapid
DNA isolations for enzymatic and hybridization
analysis. Methods Enzymol. 65:404-411.
Hall, M.N., Hereford, L., and Herskowitz, I. 1984.
Targeting of E. coli β-galactosidase to the nucleus of yeast. Cell 36:1057-1065.
Hanahan, D. 1983. Studies on transformation of
Escherichia coli with plasmids. J. Mol. Biol.
166:557-580.
Hoffman, C.S. and Winston, F. 1987. A ten-minute
DNA preparation from yeast efficiently releases
autonomous plasmids for transformation of Escherichia coli. Gene 57:267-272.
Winston, F., Chumley, F., and Fink, G.R. 1983.
Eviction and transplacement of mutant genes in
yeast. Methods Enzymol. 101:211-228.
Contributed by Charles S. Hoffman
Boston College
Chestnut Hill, Massachusetts
13.11.4
Supplement 39
Current Protocols in Molecular Biology
Preparation of Yeast RNA
UNIT 13.12
This unit provides two protocols for extraction of RNA from yeast that differ primarily
in the method for lysing the yeast cells. The first (basic) protocol isolates RNA directly
from intact yeast cells by extraction with hot acidic phenol. This procedure yields RNA
that is relatively free of contaminating DNA, is convenient to perform with multiple
samples, and gives little or no sample-to-sample variation. In contrast, the first alternate
protocol relies upon disruption of cells by vigorous mixing with glass beads and denaturing agents. Although this procedure results in efficient breaking of the cells, the product
is associated with residual DNA, and the procedure itself is troublesome when one is
working with multiple samples. A second alternate protocol describes the scaling up of
the first two procedures to isolate enough total RNA for poly (A)+ RNA preparation.
NOTE: Take precautions to avoid contamination by RNases. See
solutions, for instructions.
UNIT 4.1,
reagents and
PREPARATION OF YEAST RNA BY EXTRACTION WITH HOT
ACIDIC PHENOL
BASIC
PROTOCOL
Yeast RNA can be isolated efficiently and directly from intact cells by extraction with
acidic phenol (pH 5) and SDS at 65°C. Because this procedure does not require vortexing
individual samples with glass beads (alternate protocol), which is tedious and a source of
variability, it is well-suited for obtaining reproducible quantities of RNA from multiple
samples. In addition, RNA preparations are largely devoid of contaminating DNA which
partitions into the interface during the extraction step.
Materials
Yeast cells and desired medium (UNITS 13.1 & 13.2)
TES solution
Acid phenol
Chloroform
3 M sodium acetate, pH 5.3
100% and 70% ethanol, ice-cold
50-ml centrifuge tube (Falcon)
Centrifuge: tabletop or Sorvall equipped with an SS-34 rotor
Additional reagents and equipment for ethanol precipitation (UNIT 2.1) and
spectrophotometric quantitation of cells and RNA (APPENDIX 3)
1. Grow yeast cells in 10 ml of desired medium to mid-exponential phase (OD600 = 1.0).
It is not advisable to prepare RNA from cells that have reached a higher density because
as the stationary phase is approached, the results are less consistent and RNA yields will
vary.
2. Transfer culture to 50-ml centrifuge tube and centrifuge cells 3 min at 1500 × g (7000
rpm in a tabletop centrifuge or SS-34 rotor), 4°C.
The time and speed of the centrifugation are not critical.
3. Discard supernatant, resuspend pellet in 1 ml ice-cold water. Transfer to a clean
1.5-ml microcentrifuge tube. Microcentrifuge 10 sec at 4°C, and remove supernatant.
Proceed to step 4 or if desired immediately freeze pellet by placing tube in dry ice.
Liquid nitrogen may also be used to freeze the pellets. Although not essential, freezing is
particularly useful when RNA is to be prepared from multiple cultures or multiple time
points from a given culture; this permits simultaneous processing of the samples. The frozen
Contributed by Martine A. Collart and Salvatore Oliviero
Current Protocols in Molecular Biology (1993) 13.12.1-13.12.5
Copyright © 2000 by John Wiley & Sons, Inc.
Saccharomyces
cerevisiae
13.12.1
Supplement 23
cell pellets can be stored for months at −70°C. Thaw on ice just before continuing the
procedure.
4. Resuspend cell pellet in 400 µl TES solution. Add 400 µl acid phenol and vortex
vigorously 10 sec. Incubate 30 to 60 min at 65°C with occasional, brief vortexing.
It is crucial to incubate for ≥30 min (with occasional vortexing) to obtain quantitative
recovery of both large and small RNA species.
5. Place on ice 5 min. Microcentrifuge 5 min at top speed, 4°C.
6. Transfer aqueous (top) phase to a clean 1.5-ml microcentrifuge tube, add 400 µl acid
phenol, and vortex vigorously. Repeat step 5.
7. Transfer aqueous phase to a clean 1.5-ml microcentrifuge tube and add 400 µl
chloroform. Vortex vigorously and microcentrifuge 5 min at top speed, 4°C.
8. Transfer aqueous phase to a new tube, add 40 µl of 3 M sodium acetate, pH 5.3, and
1 ml of ice-cold 100% ethanol and precipitate. Microcentrifuge 5 min at top speed,
4°C. Wash RNA pellet by vortexing briefly in ice-cold 70% ethanol. Microcentrifuge
as before to pellet RNA.
9. Resuspend pellet in 50 µl H2O. Determine the concentration spectrophotometrically
by measuring the A260 and A280 (UNIT 4.1). Store at −70°C, or at −20°C if it is to be used
within 1 year.
Make sure that the RNA is well dissolved; if necessary, heat the resuspended pellet at 65°C
for 10 to 20 min and/or dilute further with more water. The yield from 10 ml of cells grown
in YPD medium is ∼300 ìg. Cells grown in less optimal medium will yield less RNA per ml
culture.
ALTERNATE
PROTOCOL
PREPARATION OF RNA USING GLASS BEADS
Yeast RNA is also efficiently released by disrupting the cells using high-speed mixing in
the presence of glass beads and denaturing agents. Proteins are removed by extraction
with organic solvents and the RNA is recovered by ethanol precipitation and quantitated
by measuring its absorbance at 260 nm. This preparation is suitable for S1, northern
hybridization, or primer extension analyses (UNITS 4.6, 4.9, and 4.8, respectively) and can be
prepared quickly and easily from a relatively small quantity of yeast cells. Although the
RNA isolated by this procedure is contaminated with DNA, the DNA component does
not interfere with most analytical studies.
Additional Materials
RNA buffer
25:24:1 phenol/chloroform/isoamyl alcohol (equilibrated with RNA buffer; see
support protocol, UNIT 2.1)
0.45- to 0.55-mm, chilled, acid-washed glass beads (Sigma)
Prepare the cells
1. Grow and process yeast cells and freeze cell pellet as in steps 1 to 3 of the basic
protocol.
If necessary, the samples can now be quick-frozen on dry ice and stored at −70°C. Thaw
on ice just before processing.
2. Resuspend pellet in 300 µl RNA buffer.
Preparation of
Yeast RNA
Disrupt the cells
3. Add a volume of chilled acid-washed glass beads equivalent to ∼200 µl water.
13.12.2
Supplement 23
Current Protocols in Molecular Biology
Prepare acid-washed glass beads by soaking in concentrated nitric acid for 1 hr, washing
extensively with deionized water, and drying in a baking oven.
Before use the glass beads should be chilled on ice. Manipulate the beads with a stainless
steel spatula that has been dried in a baking oven. Do this carefully, using a spatula or a
funnel fashioned out of weighing paper. If beads stick to the lip of the microcentrifuge tube,
they will prevent the tube from closing securely.
4. Add 300 µl of 25:24:1 phenol/chloroform/isoamyl alcohol equilibrated with RNA
buffer.
A phenol mix previously equilibrated with TE buffer (UNIT 2.1) can be reequilibrated with
RNA buffer. Remove the aqueous layer (TE buffer), add a volume of RNA buffer equal to
the volume of the organic layer, shake vigorously, and allow the phases to separate. Remove
the aqueous layer (RNA buffer), add fresh RNA buffer, mix well, and allow the phases to
separate.
5. Close the cap, then invert and shake up and down to ensure that the beads are
suspended. Vortex vigorously for 2 min at highest speed.
Hold 2 to 4 tubes on the head of a vortexer for 1 min, place these tubes on ice, and vortex
another set. After 1 min with the second set, place on ice, and vortex the first set for another
minute. This limits the processing to 8 samples at one time. If a high-speed horizontal shaker
is available many samples can be vortexed simultaneously. Use the highest speed setting
for 3 min in a cold room.
6. Microcentrifuge 1 min at room temperature. Transfer aqueous (top) layer to a clean
microcentrifuge tube.
Avoid the interface by taking only the uppermost 200 to 250 ìl from each sample. Taking
a fixed amount from each sample results in a more uniform yield of RNA from each sample.
7. Add an equal volume of 25:24:1 phenol/chloroform/isoamyl alcohol (200 to 250 µl).
Vortex vigorously 10 sec.
8. Repeat step 6.
Precipitate the RNA
9. Add 3 vol (∼600 µl) of ice-cold 100% ethanol. Mix well and place at −20°C for ≥30
min or on dry ice for 5 min.
10. Microcentrifuge 2 min at 4°C. Aspirate or pour off the supernatant and wash pellet
with ice-cold 70% ethanol.
11. Microcentrifuge 1 min at 4°C. Aspirate or pour off supernatant and dry pellet.
12. Resuspend pellet in 50 µl H2O. Determine the concentration spectrophotometrically
by measuring the A260 and A280 (UNIT 4.1). Store the RNA at −70°C.
If 2 × 108 cells are used, the RNA concentration of the final solution will be ∼2 mg/ml.
Saccharomyces
cerevisiae
13.12.3
Current Protocols in Molecular Biology
Supplement 23
ALTERNATE
PROTOCOL
PREPARATION OF POLY(A)+ RNA
The RNA isolated in the basic and first alternate protocols is also a suitable source of
poly(A)+ RNA for use in constructing cDNA libraries. For this purpose, larger quantities
of RNA can be isolated by simply scaling-up the procedures. For 1010 S. cerevisiae cells,
use the following guidelines:
For the hot acidic phenol protocol: Increase volumes of TES and acid phenol solutions
to 4 ml each; use 50-ml polypropylene centrifuge tubes for all manipulations. The yield
will be ∼10 mg of RNA.
For the glass beads protocol: Increase volumes to 15 ml RNA buffer, 10-ml volume glass
beads, 15 ml phenol/chloroform/isoamyl alcohol, and 30 ml of 100% ethanol. Perform
the procedure in a 50-ml disposable polypropylene tube and centrifuge 10 min, 3000 rpm
(1200 × g), 4°C in a fixed-angle or swinging-bucket type rotor at each phenol extraction
step. Good recovery of the precipitated nucleic acid can be accomplished by centrifugation under these same conditions. The yield will be ∼5 mg total RNA.
Prepare poly(A)+ RNA from either method using the protocol presented in UNIT 4.5.
REAGENTS AND SOLUTIONS
Acid phenol
Add sufficient water to a bottle of solid phenol such that phenol is water-saturated;
pH will be ∼5.0. Do not buffer phenol. Store at 4°C, protected from light.
RNA buffer
0.5 M NaCl
200 mM Tris⋅Cl, pH 7.5
10 mM EDTA
Store indefinitely at room temperature
TES solution
10 mM Tris⋅Cl, pH 7.5
10 mM EDTA
0.5% SDS
Store indefinitely at room temperature
COMMENTARY
Background Information
Preparation of
Yeast RNA
Aside from the harsh conditions used to
break open yeast cells, the RNA isolation procedures presented here are similar to the phenol/SDS method for isolating RNA from plant
cells (UNIT 4.3). Both methods take advantage of
the fact that phenol extraction is an effective
means of inactivating and removing RNases.
The hot acidic phenol method is preferable
when working with multiple samples; because
the number of manipulations and time required
for each are reduced, very little sample-to-sample variation is encountered.
The glass bead disruption procedure causes
the release of both RNA and DNA, and the
absorbance at 260 nm will measure total nucleic
acid. For precise determination of RNA con-
centrations, the DNA component can be removed as described in UNITS 4.1, 4.3, and 4.5. This
is not necessary for routine northern blot,
primer extension, and S1 analyses. Refer to
UNITS 4.1-4.5 for further information on the purification and properties of RNA.
Critical Parameters
As with all RNA manipulations, precautions
must be taken to prevent RNase contamination.
See UNIT 4.1 for details. In the hot acidic phenol
method, the two phenol and one chloroform
extractions are critical to obtain clean RNA for
analysis.
These protocols are designed to process
multiple samples in tandem—for example,
when monitoring gene induction over time or
13.12.4
Supplement 23
Current Protocols in Molecular Biology
when examining RNA synthesis driven by different promoter constructions. In these cases
(where quantitation is critical), detection using
probes in northern hybridization, S1 mapping,
or primer extension studies should be carried
out in parallel (preferably in the same tube or
hybridization bag) with a probe for a gene in
which transcription does not change over time
or in response to inducing conditions. The ratios of signal intensities for the two probes in
each sample are then compared.
Anticipated Results
The yield of RNA isolated using the hot
acidic phenol protocol is ∼300 µg from 2 × 108.
The yield resulting from the glass bead disruption protocol starting with 2 × 108 cells will be
∼100 µg of total RNA. Scale-up of either
method will produce ∼5 to 10 mg total RNA.
Ten to twenty micrograms of total RNA isolated by either procedure is sufficient for most
northern blot, S1 mapping, or primer extension
applications.
Time Considerations
Depending on the organization of the researcher, 12 to 24 samples can be processed
conveniently in about 1 hr using the hot acidic
phenol method. For the glass bead method,
eight samples can be easily processed without
using a high-speed shaking apparatus in ∼1.5
hr. A second set of eight samples can be processed while the first set is precipitating at −
20°C. With a high-speed shaking apparatus 16
samples can be processed in about 1 hr. Both
phenol extraction steps can be done on the
shaking apparatus.
If RNA synthesis is being monitored over
time, samples from individual time points can
be frozen at ∼70°C (see step 3, basic protocol)
so that all of the samples can be processed
simultaneously.
Contributed by Martine A. Collart and
Salvatore Oliviero
Harvard Medical School
Boston, Massachusetts
Saccharomyces
cerevisiae
13.12.5
Current Protocols in Molecular Biology
Supplement 23
UNIT 13.13
Preparation of Protein Extracts from Yeast
Three protocols are presented for preparing protein extracts; they differ primarily in the
way the cells are broken. In the basic protocol, cells are enzymatically converted to
spheroplasts, which are then lysed by a combination of osmotic shock and Dounce
homogenization. A support protocol for isolating intact nuclei by differential
centrifugation is also presented. An alternate protocol describes mechanical breakage of
cells by vortexing in the presence of glass beads. In a second alternate protocol, growing
cells are frozen immediately in liquid nitrogen and then lysed by grinding in an industrial-strength blender in the presence of liquid nitrogen. These methods each have
advantages and disadvantages; the best choice will depend on the particular application.
For all of these procedures, it is advantageous to use protease-deficient strains such as
BJ926 or EJ101.
BASIC
PROTOCOL
SPHEROPLAST PREPARATION AND LYSIS
Materials
Protease-deficient yeast cells (BJ926, EJ101, or equivalent)
YPD medium (UNIT 13.1)
Zymolyase buffer, room temperature and ice-cold
Zymolyase 100T (ICN Immunobiologicals)
1 M sorbitol (optional)
Lysis buffer
Extraction buffer
Storage buffer
Liquid nitrogen
Sorvall GS-3 or GSA rotor (or equivalent)
Sorvall SS-34 or SA-600 rotor (or equivalent)
Beckman Type 45Ti rotor (or equivalent; large capacity)
30°C shaker platform
Rubber policeman
Dounce homogenizer
Rotating wheel or rocker
Additional reagents and equipment for growing yeast cells (UNITS 13.1 & 13.2),
large-volume dialysis (APPENDIX 3), and determining conductivity (UNIT 10.10)
Grow the cells
1. Grow cells to mid-log phase in YPD medium with vigorous shaking or forced
aeration.
The OD600 can vary with aeration conditions, but should be between 1 and 5. The procedure
is appropriate for between 100-ml and 20-liter cultures, although it can be scaled up if
necessary.
2. Harvest cells by centrifugation 5 min at 1500 × g (GS-3 or GSA rotor at ∼3000 rpm),
4°C, in preweighed centrifuge bottles.
Prepare the spheroplasts
3. Determine the wet weight (in grams) of yeast cells in the pellet by the weight increase
over that of the preweighed bottle. This is approximately equal to the packed cell
volume (in milliliters), and for all subsequent steps will be considered 1 vol.
Preparation of
Protein Extracts
from Yeast
13.13.1
Supplement 23
One liter of BJ926 (a diploid strain) at OD600 = 1.0 yields a packed cell volume of 2 to 3
ml.
Contributed by Barbara Dunn and C. Richard Wobbe
Current Protocols in Molecular Biology (1992) 13.3.1-13.3.9
Copyright © 2000 by John Wiley & Sons, Inc.
4. Resuspend cells in 2 to 4 vol ice-cold water and immediately centrifuge 5 min at 1500
× g (SS-34 or SA-600 rotor at 3500 rpm), 4°C. Discard the supernatant.
5. Resuspend the cells by adding 1 vol zymolyase buffer containing 30 mM DTT (see
note in reagents and solutions), and incubate 15 min at room temperature.
This step facilitates subsequent zymolyase treatment and spheroplast lysis by breaking
disulfide bonds.
6. Centrifuge 5 min at 1500 × g, 4°C, and resuspend in 3 vol zymolyase buffer. Add 2
mg (200 U) Zymolyase 100T per ml of original packed cell volume to the resuspended
cells. Incubate 40 min at 30°C on a shaker platform at ∼50 rpm.
7. Determine if conversion to spheroplasts has been completed by the lysis in water
technique (UNIT 13.7). If spheroplasting is incomplete, continue incubation until
complete.
Perform all procedures from this point on at 4°C.
8. Centrifuge spheroplasts 5 min at 1500 × g. Decant the supernatant carefully—the
spheroplast pellet will not be as tight as the previous cell pellets.
For some procedures (e.g., extracts for in vitro transcription or translation), spheroplasts
are resuspended in YPD medium containing 1 M sorbitol and incubated for 30 to 60 min
at 30°C to allow metabolic recovery.
9. Wash the spheroplasts by gently resuspending the pellet in 2 vol ice-cold zymolyase
buffer and centrifuging 5 min at 1500 × g. Repeat this step two more times.
Spheroplasts are sticky and difficult to resuspend. To facilitate resuspension, first resuspend
the spheroplasts in a small volume with the aid of a rubber policeman and then add more
buffer to achieve the correct final volume.
The washing step is important for removing proteases, phosphatases, and nucleases present
in the zymolyase preparation. For some purposes, it may be necessary to carry out
additional washes.
Lyse the spheroplasts
10. Gently resuspend the pellet in 2 vol lysis buffer. Do not try to achieve a homogeneous
suspension; simply dislodge the pellet from the side of the centrifuge tube and gently
swirl 10 to 20 times. Centrifuge spheroplasts 10 min at 1500 × g.
Extensive manipulation of the pellet may result in premature osmotic lysis.
11. Thoroughly resuspend spheroplast pellet with 1 vol lysis buffer using a glass rod.
At this point, tubes containing the resuspended spheroplasts can be quick-frozen in liquid
nitrogen and stored at −80°C. Thaw frozen spheroplasts overnight on ice before proceeding.
12. Lyse spheroplasts with 15 to 20 strokes of a tight-fitting pestle (clearance 1 to 3 µm)
in a Dounce homogenizer.
Extract proteins from lysate
13. Half-fill ultracentrifuge tubes with lysate. Add an equal volume of extraction buffer
and seal the tubes. Gently invert tubes on rotating wheel or rocker for 15 to 30 min
at 4°C.
The lysate and extraction buffer are not premixed because the resulting solution will
immediately become quite viscous and hence difficult to pour if the ionic strength is above
∼0.5 M.
Saccharomyces
cerevisiae
13.13.2
Current Protocols in Molecular Biology
Supplement 12
14. Centrifuge 90 min at 100,000 × g (Type 45Ti rotor at 33,000 rpm), 4°C.
15. Collect supernatant and dialyze 2 to 4 hr against 100 vol storage buffer. Transfer
dialysis bag to 100 vol fresh storage buffer; dialyze an additional 2 to 4 hr.
A flocculent precipitate may form during dialysis. These precipitates usually contain
negligible amounts of most protein factors and can be discarded.
16. Remove a few microliters of the dialysate, dilute 1:1000 with water, and determine
the conductivity (UNIT 10.10). If it is equal to that of similarly diluted storage buffer or
below some acceptable value (usually 100 to 250 mM NaCl), proceed to step 17. If
not, continue dialysis.
17. Centrifuge dialysate 10 min at 10,000 × g (SS-34 rotor at 9200 rpm or SA-600 rotor
at 8500 rpm), 4°C. Collect the supernatant, freeze in small aliquots in liquid nitrogen,
and store at −80°C.
This crude extract contains most DNA-binding proteins as well as transcription and
replication factors. The pellet contains proteins that can be “salted in” by resuspending in
storage buffer containing 0.5 to 1.0 M KCl, if desired.
SUPPORT
PROTOCOL
NUCLEI PREPARATION BY DIFFERENTIAL CENTRIFUGATION
Nuclei suitable for chromatin studies and/or nuclear protein extracts are prepared by
osmotically lysing spheroplasts in the presence of Ficoll, which preserves nuclear
structure and prevents proteins from leaking out of the nucleus, followed by differential
centrifugation.
Additional Materials
Ficoll buffer, ice-cold
Teflon pestle tissue homogenizer, motor-driven (optional; Thomas)
1. Perform steps 1 to 9 of basic protocol. Resuspend cells in 0.5 vol zymolyase buffer.
2. Pipet cells drop by drop into a beaker containing 15 to 25 vol ice-cold Ficoll buffer
with continuous stirring in an ice bath or cold room.
Alternatively, resuspend cells in several volumes Ficoll buffer and homogenize with 5 to
10 strokes using a motor-driven tissue homogenizer and moderately tight Teflon pestle
(clearance 0.15 to 0.23 mm) at medium speed.
3. Transfer the suspension to centrifuge tubes and centrifuge 5 min at 3000 × g (SS-34
rotor at 5000 rpm), 4°C, to pellet cell debris and unlysed spheroplasts.
This step can be skipped if separation of unlysed cells is not critical, or it can be repeated
multiple times if it is important to completely remove debris.
4. Transfer supernatant to new centrifuge tubes and centrifuge 20 min at 20,000 × g
(SS-34 rotor at 13,000 rpm), 4°C. Decant the supernatant. The pellet contains the
nuclei.
The pellet can be difficult to resuspend. If necessary, resuspend as described in step 9 of
basic protocol.
Preparation of
Protein Extracts
from Yeast
For chromatin studies, the nuclei can be suspended in an appropriate buffer and used
directly for micrococcal nuclease or DNase I digestion. In some instances, it is useful to
wash the nuclei in a buffer equivalent to Ficoll buffer but containing 10% glycerol instead
of Ficoll. Nuclei can be stored in this buffer several months at −80°C if the tubes are frozen
quickly on dry ice or liquid nitrogen.
5. To obtain nuclear extracts, resuspend the nuclei in 1 vol lysis buffer and perform
13.13.3
Supplement 12
Current Protocols in Molecular Biology
steps 12 to 17 of basic protocol.
Alternatively, the nuclei can be washed in buffers containing various types and molarities
of salt. The supernatants will contain proteins that have been extracted by the salt.
CELL DISRUPTION USING GLASS BEADS
ALTERNATE
PROTOCOL
These procedures are for small-scale preparations using a vortexer, and for large-scale
preparations using a Bead Beater.
Additional Materials
Glass bead disruption buffer
Chilled, acid-washed glass beads (0.45- to 0.55-mm; see reagents and solutions)
Bead Beater and vessel (Biospec; optional)
1. Grow and harvest yeast cells as described in steps 1 to 4 of the basic protocol for
spheroplast lysis. Determine the packed cell volume.
All subsequent steps should be carried out at 4°C.
2. Resuspend cells in 1 vol glass bead disruption buffer.
The cells can be frozen by slowly pouring this suspension into a plastic beaker filled with
liquid nitrogen. Use enough liquid nitrogen to submerge the frozen paste. This frozen
“popcorn” can be stored at −80°C. Thaw overnight on ice before proceeding.
3. Mix cell paste with 2 vol glass bead disruption buffer.
4. Add 4 vol of chilled, acid-washed glass beads.
For packed cell volumes of <10 ml:
5a. Transfer the cell suspension to an appropriately sized screw-cap centrifuge tube. The
suspension should occupy no more than 60% to 70% of the capacity of the tube.
6a. Vortex the suspension at maximum speed for 30 to 60 sec at 4°C; place tube on ice
for 1 to 2 min. Repeat 3 to 5 more times. Check the amount of cell breakage by visual
inspection under a microscope. Proceed to step 7.
Although larger volumes may be processed by using multiple aliquots, it is better to use a
Bead Beater as described below.
For packed cell volumes of >10 to 20 ml:
5b. Transfer the cell suspension to an appropriately sized Bead Beater vessel (stainless
steel is recommended for better heat transfer) and add glass bead disruption buffer
to fill the vessel almost to the brim. The volume of buffer required to fill the vessel
should not exceed 1 cell-suspension volume. Attach the blade and cap assembly,
ensuring that all air is excluded from the vessel.
It is important to exclude air in order to prevent foaming and potential protein denaturation.
6b. Grind at high speed for 60 sec, then let sit 1 to 2 min on ice. Repeat 3 to 5 more times.
7. Allow the glass beads to settle out and decant the supernatant.
8. Add 2 to 4 vol glass bead disruption buffer to the glass beads and invert the tube 5 to
10 times. Allow the beads to resettle and decant the supernatant. Pool the supernatants.
9. Centrifuge the pooled supernatants 60 min at 12,000 × g (SS-34 rotor at 10,000 rpm),
Saccharomyces
cerevisiae
13.13.4
Current Protocols in Molecular Biology
Supplement 12
4°C. Collect the supernatant which represents the crude extract. For long-term
storage, aliquot into small tubes, quick-freeze the tubes in liquid nitrogen, and store
at −80°C.
ALTERNATE
PROTOCOL
CELL DISRUPTION USING LIQUID NITROGEN
This protocol is designed for processing 200 ml to ∼20 liters of cells. It can be easily
scaled up for processing larger amounts (e.g., from a fermentor), in which case a larger
blender and cup must be used.
Additional Materials
Yeast cakes (optional; Red Star)
Liquid nitrogen
60-ml syringe
1-liter plastic beaker (Nalgene or equivalent)
1-liter stainless steel blender cup and blender (Waring or equivalent)
1. Grow cells to mid-log phase in YPD (or selective) medium with vigorous shaking or
forced aeration. Centrifuge the culture to harvest cells. Discard supernatant.
Many liters can be processed at one time with this method. Alternatively, commercial yeast
cakes can be used in place of the growing cells.
2. Vortex cell pellet to create a thick cell paste. If necessary, add a minimal amount of
ice-cold water to allow the paste to be poured or spooned.
NOTE: Keep cell paste on ice in subsequent steps.
If using a yeast cake, mix equal volumes of cells and water and blend into a thick paste.
3. Remove the plunger from a 60-ml syringe. Seal the bottom with a plug or tightly
wrapped parafilm. Pour or spoon in 50 ml of the cell paste.
4. Place 400 ml liquid nitrogen into a 1-liter Nalgene beaker.
Avoid contact with liquid nitrogen due to the danger of frostbite. Use insulated gloves when
handling containers. Never use glass containers to hold liquid nitrogen as they may break.
5. Hold the syringe over the liquid nitrogen; remove plug or parafilm from bottom, insert
plunger, and squeeze the cell paste into the liquid nitrogen. Repeat steps 3 and 5 no
more than three times. A maximum of ∼200 ml cell paste can be processed at one
time when using a 1-liter blender cup.
Long spaghetti-like aggregations of frozen cells should form with a thick cell paste;
spherical popcorn-like clumps form with a thinner paste. The frozen cells can be stored
indefinitely at −80°C before disruption.
6. Carefully pour the liquid nitrogen with frozen cells into a blender cup that is already
attached to the blender and has been thoroughly dried prior to use.
It is preferable to perform the grinding in a cold room with a precooled blender to decrease
the rate of evaporation of the liquid nitrogen, but the procedure can be done at room
temperature if necessary.
Preparation of
Protein Extracts
from Yeast
Only industrial-strength blenders and stainless steel blender cups can be used for this
protocol (a Waring commercial blender is recommended). Care should be taken that all
moving parts of both the blender and blender cup are completely free of moisture in order
to avoid freezing when adding the liquid nitrogen.
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7. Place the lid on the blender cup, making sure the lid vents are open so that pressure
does not build up in cup as liquid nitrogen evaporates. Keeping the lid held down
tightly, grind at high speed in three successive 2-min bursts. Between bursts, mix
frozen powder and, if necessary, add liquid nitrogen to just cover powder.
Begin grinding as soon as possible after pouring in the liquid nitrogen to avoid freezing
the moving parts. Some material will spurt out of the lid vents during grinding; take care
not to get in its way. It is advisable to cover the working area with plastic sheeting prior
to grinding to ease the clean-up. Pulsed grinding may help avoid the spurting problem, but
keep in mind that speed is critical here—the frozen powder should not be allowed to thaw.
8. After grinding, pour the fine frozen yeast powder into a beaker containing twice the
original cell-paste volume of ice-cold storage buffer. Mix and then pour the suspension into a centrifuge bottle. Keep bottle on ice.
9. Repeat steps 3 through 8 until all the cell paste has been processed.
10. Centrifuge 15 min at 5000 × g (GSA or GS-3 rotor at 5500 rpm; or SS-34 rotor at
6500 rpm), 4°C. The supernatant contains a crude extract whose concentration should
be between 10 and 20 mg protein/ml. It can be used immediately for assays or as a
starting point for protein purification. For long-term storage, aliquot the supernatant
into plastic tubes, freeze quickly on dry ice, and store at −80°C.
Alternatively, the frozen yeast powder can be resuspended in 1 vol lysis buffer and treated
as described in steps 13 to 17 of the basic protocol for spheroplast lysis.
REAGENTS AND SOLUTIONS
Extraction buffer
Lysis buffer (see below)
0.8 M ammonium sulfate
20% glycerol
Ficoll buffer
18% (w/v) Ficoll-400 (Pharmacia)
10 mM Tris⋅Cl, pH 7.5
20 mM KCl
5 mM MgCl2
3 mM DTT
1 mM EDTA
1× protease inhibitor mix (see below)
1 mM PMSF
Glass bead disruption buffer
20 mM Tris⋅Cl, pH 7.9
10 mM MgCl2
1 mM EDTA
5% glycerol
1 mM DTT
0.3 M ammonium sulfate
1× protease inhibitor mix (see below)
1 mM PMSF
The concentration of ammonium sulfate in the buffer can be varied between 0.1 and 1.0 M.
Final concentrations above 0.25 M strip specific DNA-binding proteins and histones off
chromatin and are therefore useful in obtaining factors that interact with nucleic acids. KCl
and NaCl can also be added to final concentrations between 0.1 and 2.0 M.
Saccharomyces
cerevisiae
13.13.6
Current Protocols in Molecular Biology
Supplement 24
Glass beads, chilled and acid-washed, 0.45- to 0.55-mm
Wash the beads by soaking 1 hr in concentrated nitric acid. Rinse thoroughly with water.
Dry the beads in a baking oven, cool to room temperature, and store at 4°C until needed.
Lysis buffer
50 mM Tris⋅Cl, pH 7.5
10 mM MgSO4
1 mM EDTA
10 mM potassium acetate
1 mM DTT
1× protease inhibitor mix (see below)
1 mM phenylmethylsulfonyl fluoride (PMSF)
100× protease inhibitor mix
Listed below are representative protease inhibitors; different combinations may be
more appropriate for individual applications.
10 µg/ml chymostatin
200 µg/ml aprotinin
100 µg/ml pepstatin A
110 µg/ml phosphoramidon
720 µg/ml E-64
50 µg/ml leupeptin
250 µg/ml antipain
10 mM benzamidine
10 mM sodium metabisulfite
Storage buffer
20 mM Tris⋅Cl, pH 7.5
0.1 mM EDTA
10% glycerol
100 mM KCl
1 mM DTT
1× protease inhibitor mix (see above)
1 mM PMSF
Zymolyase buffer
50 mM Tris⋅Cl, pH 7.5
10 mM MgCl2
1 M sorbitol
1 mM or 30 mM DTT (see annotation below)
For step 5 of basic protocol use 30 mM DTT in this buffer. For all other applications, use 1
mM DTT.
COMMENTARY
Background Information
Preparation of
Protein Extracts
from Yeast
The three protocols for preparing protein
extracts from yeast differ primarily in the way
the cells are broken. Each of these methods has advantages and disadvantages that
should be considered when choosing one for
a particular application. Most importantly,
the extracts made by these different methods
may contain different proteins and different
levels of protein activity. Thus, it may be
necessary to try different methods to obtain
extracts capable of carrying out the in vitro
reactions of interest.
Spheroplast procedure. Cells are converted
to spheroplasts by zymolyase treatment, and
then lysed by a combination of osmotic shock
and Dounce homogenization. Because this
method is the most gentle way to break yeast
cells, it is most suitable for preparing extracts that can carry out complex enzymatic
13.13.7
Supplement 24
Current Protocols in Molecular Biology
functions (e.g., translation, transcription, DNA
replication) and in which the integrity of
macromolecular structures (e.g., ribosomes,
splicesomes) has been maintained. It is also
useful for isolating intact nuclei that can be used
for chromatin studies (Bloom and Carbon,
1982) or for nuclear protein extracts (Lue and
Kornberg, 1987). The support protocol described here for isolating nuclei derives primarily from Nelson and Fangman (1979) and is
based on differential centrifugation in the presence of high concentrations of Ficoll (a polymer that inhibits leakage of nuclear proteins
into the cytoplasm). Another procedure for purifying the nuclei (after release by spheroplast
lysis) employs a Percoll gradient (Amati and
Gasser, 1988). Although these nuclei are
slightly more purified, they seem to be of varying or lower quality than those produced by the
differential centrifugation (based on nucleosome ladder integrity). The major disadvantages of the spheroplast lysis procedure are
that it is relatively tedious and expensive, especially for large-scale preparations (>10 liters),
and the long incubation periods can lead to
proteolysis or protein modification.
Liquid nitrogen procedure. Initially described by Sorger and Pelham (1987), cells are
frozen immediately in liquid nitrogen and then
lysed by grinding in a Waring blender in the
presence of liquid nitrogen. The protocol is
quick and easy (albeit a bit messy and potentially dangerous to the careless investigator). It
can accommodate varying amounts of yeast
cells including very large cultures. Its main
advantage is that cells are taken immediately
from the actively growing state into liquid nitrogen (−135°C), decreasing degradative enzyme activities such as proteases and nucleases
as well as activities that modify proteins (e.g.,
phosphatases and kinases). It is particularly
suited for making whole-cell extracts from a
single yeast culture for large-scale protein purification. A detailed description of the technique as applied to very large-scale cultures
(100 to 400 liters) is given by Sorger et al.
(1989). A drawback to the liquid nitrogen protocol is that small samples (i.e., 10- to 100-ml
yeast cultures) are not easily processed because
there is not enough mass of frozen cell clumps
to fracture effectively in the blender. Stainless
steel blender cups with very small capacities
(10 to 100 ml) can be obtained and may be
useful for processing such small amounts. Multiple samples pose another problem for the
liquid nitrogen technique because, usually,
only one blender is available, making it time-
consuming to process individual samples and
to clean the equipment between uses.
Glass bead procedure. This protocol involves mechanically breaking cells by vortexing in the presence of glass beads. It is very
flexible because it can be carried out easily on
multiple small (10-ml) cultures as well as on
very large cultures with appropriate equipment
such as a Bead Beater (Klekamp and Weil,
1982). It is particularly useful when making
extracts from many different small yeast cultures for assaying purposes rather than for protein purification. Unfortunately, during the
glass bead procedure, proteins are treated
harshly causing extensive foaming. The
amount of cell breakage varies, and proteolysis
and modification of the proteins may result
from heating the extract above 4°C during the
mechanical breakage.
Critical Parameters
For all these procedures, it is advantageous
to use protease-deficient strains such as BJ926
or EJ101, but laboratory strains of a desired
genotype can be used. In addition, commercial
yeast cakes can be used for some applications
and have the advantage that limitless quantities
of material can be obtained for little cost and
no work. The liquid nitrogen method is probably best for processing yeast cakes, while the
spheroplast method is unsuitable for this application.
Another crucial factor, even for extracts
from protease-deficient strains, is the inclusion
of a wide collection of protease inhibitors at all
stages during and after lysis. The protease inhibitor mix given in this procedure, along with
PMSF and EDTA (to inhibit metalloproteases)
should inactivate almost all types of proteases,
although other combinations may be more appropriate for particular applications. A final
general consideration is to be sure that all solutions coming in contact with the extracted
proteins are kept ice cold.
Spheroplast procedure. For this protocol, it
is important to handle the spheroplasts gently.
The amount of time and zymolyase needed to
achieve complete spheroplasting can vary quite
a bit between strains, and some adjustments of
the amounts given in the protocol may be necessary. Complete spheroplasting is essential for
efficient lysis. Only the highest grade Zymolyase 100T should be used. However, because even this grade of zymolyase contains
considerable proteolytic activity, it is important
to wash the spheroplasts several times prior to
lysis.
Saccharomyces
cerevisiae
13.13.8
Current Protocols in Molecular Biology
Supplement 13
Liquid nitrogen procedure. For optimal
yields of protein, all (or nearly all) the cells
should be broken. This should be tested by
suspending a small amount of the ground powder in buffer and visually inspecting under the
microscope. (Remember to add more liquid
nitrogen to the remaining powder to keep it
frozen.) If many intact (phase-refractile) cells
are seen, further grinding in liquid nitrogen
should be performed. Care should be taken to
ensure that the liquid nitrogen covering the cells
does not evaporate. The frozen yeast powder
should always be poured into cold extraction
buffer as quickly as possible.
Glass bead procedure. It is crucial that the
beads be the correct size. Cell breakage will be
inefficient if the wrong size beads are used. As
is the case for the liquid nitrogen procedure, the
degree of breakage should be examined before
proceeding. Finally, it is very important that the
temperature of the solution during the breakage
remains between 0° and 4°C. Because mechanical disruption generates heat, it is usually
necessary to vortex in short bursts so that the
solution cools down between intervals. It is also
important to minimize foaming during vortexing as this is indicative of protein denaturation
which may result in considerable loss of activity. Because the Bead Beater generates much
less foaming than vortexing, it is preferred for
large-scale preparations.
Anticipated Results
Cell extracts produced by any of these procedures contain ∼10 to 30 mg/ml of the total
cellular protein. These extracts can be used
directly for some purposes, such as mobility
shift DNA binding assays (Chapter 12), or can
serve as the first step in protein purification
(Chapter 10). The extracts will also contain
large amounts of nucleic acid, especially RNA.
Nuclear extracts will contain significant concentrations of chromosomal DNA.
Time Considerations
ing on the amount of yeast cells being processed. The most time-consuming and tedious
steps are the resuspensions of the spheroplasts.
Isolation of nuclei will take an additional 2 to
4 hr depending on the size of the preparation.
For the liquid nitrogen or glass bead procedures, several liters of a single culture sample
can be processed in an hour or less; larger
cultures require more time.
Literature Cited
Amati, B.B. and Gasser, S.M. 1988. Chromosomal
ARS and CEN elements bind specifically to the
yeast nuclear scaffold. Cell 54:967-978.
Bloom, K.S. and Carbon, J. 1982. Yeast centromere
DNA is in a unique and highly ordered structure
in chromosomes and small circular minichromosomes. Cell 29:305-317.
Klekamp, M.S. and Weil, P.A. 1982. Specific transcription of homologous class III genes in yeast
Saccharomyces cerevisiae soluble cell-free extracts. J. Biol. Chem. 257:8432-8441.
Lue, N.F. and Kornberg, R.D. 1987. Accurate initiation at RNA polymerase II promoters in extracts
from Saccharomyces cerevisiae. Proc. Natl.
Acad. Sci. U.S.A. 84:8839-8843.
Nelson, R.G. and Fangman, W.L. 1979. Nucleosome organization of the yeast 2µ DNA plasmid: A eukaryotic minichromosome. Proc. Natl.
Acad. Sci. U.S.A. 76:6515-6519.
Sorger, P.K. and Pelham, H.R.B. 1987. Purification
and characterization of a heat-shock element
binding protein from yeast. EMBO J. 6:30353041.
Sorger, P.K., Ammerer, G., and Shore, D. 1989.
Identification and purification of sequence-specific DNA-binding proteins. In Protein Function:
A Practical Approach (T.E. Creighton, ed.) pp.
199-223. IRL Press. Oxford.
Contributed by Barbara Dunn
Memorial Sloan-Kettering Cancer Center
New York, New York
C. Richard Wobbe
Harvard Medical School
Boston, Massachusetts
After growing the cells, the spheroplast procedure should take between 3 to 12 hr depend-
Preparation of
Protein Extracts
from Yeast
13.13.9
Supplement 13
Current Protocols in Molecular Biology
SCHIZOSACCHAROMYCES POMBE
SECTION V
Overview of Schizosaccharomyces pombe
UNIT 13.14
The fission yeast S. pombe is a popular
model eukaryote that offers complementary
strengths to the more familiar budding yeast S.
cerevisiae. The organisms are evolutionarily
distant with an estimated 1000 million years
separation. Fission yeast has been dubbed an
archaeascomycete, making it quite separate
from common ascomycetes, including S. cerevisiae and the filamentous fungi (Hedges,
2002); however, like budding yeast, fission
yeast is genetically tractable, and lends itself to
easy molecular manipulation. Its increasing
popularity in recent years, especially in labs
studying mammalian cell biology, has led to the
nickname “micromammal.”
Fission yeast was discovered in 1893, but
wasn’t commonly taken up as a model organism until the 1960s (e.g., see Mitchison, 1990;
Egel, 2000; Yanagida, 2002), largely for studies
of growth control and differentiation. In the
1970s, Nobel laureate Paul Nurse and his colleagues isolated cell cycle mutants that either
blocked the cell division cycle, or accelerated
it, making S. pombe an important model for cell
cycle control (Nurse, 2000). S. pombe continues as an important system for studying cell
cycle regulation, sexual differentiation, and
more recently, chromosome dynamics and polarized cell growth. Other areas were less developed but are now growing as new investigators continue to move into the system. Given
its wide divergence from S. cerevisiae, S.
pombe provides a complementary model system, and the compare and contrast approach to
problems using both species has proven remarkably fruitful in providing models for biology of larger cell types (e.g., Forsburg, 1999).
With a haploid genome size of just 13.8 Mb,
distributed amongst 4,824 open reading frames
(compared to over 5500 for S. cerevisiae), S.
pombe is the smallest sequenced eukaryote
(Wood et al., 2002). Unlike S. cerevisiae, there
is no evidence of genome-wide duplications in
S. pombe.
There are ∼145 genes in S. pombe that have
metazoan homologs but are not found in S.
cerevisiae, and a similar number vice versa
(Wood et al., 2002). These genes tend to represent groups of proteins involved in particular
cellular functions. For example, S. pombe has
three relatively large chromosomes of 5.7, 4.6,
and 3.5 Mb. These have features typical of
higher eukaryotes, including large diffuse replication origins (∼1 kb) and large heterochromatic centromeres (∼40 to 100 kb). These elements do not contain short consensus sequences, as in the more numerous small
chromosomes of budding yeast, but are defined
by activity typical of the same elements in
higher cells. Many of the proteins common to
S. pombe and metazoans that are missing in S.
cerevisiae are implicated in chromosome functions, including heterochromatin factors such
as Swi6/HP1, chromatin modifiers such as the
histone methyltransferase Clr4/SuVar2-9,
telomere proteins such as Taz1/TRF, centromere proteins such as CENP-B, and components of RNAi apparatus such as Argonaut and
Dicer. Not surprisingly, S. pombe is proving to
be a particularly appropriate model system for
studies of mammalian chromosome biology.
Similarly, given its distinct rod-shaped morphology and division by medial fission, it is
also a good model for polarity and cytokinesis.
Other problems are also easily approached.
The choice of system is in part driven by the
problem under study, but also by the availability
of local expertise. Relatively few U.S. investigators chose fission yeast as a system prior to
the explosion of the cell cycle studies in the
1980s, in contrast to its strong popularity in
Japan and Europe; however, S. pombe is now
well represented throughout North America, as
well as in its traditional strongholds. In addition, a number of groups with experience in
mammalian cell biology have picked up S.
pombe as their alternative model system, leading to a substantial group of fission yeast researchers who work in labs that do not consider
themselves yeast groups. The fission yeast
community overall is relatively small by comparison to that of budding yeast but is friendly
and international in outlook. It is held together
by an international meeting in alternate years,
and extensive online resources.
There is a persistent myth suggesting that S.
pombe cannot be manipulated to the same extent as budding yeast. In practice, working in
S. pombe is not strikingly different from working in S. cerevisiae. Most tools available in S.
Yeast
Contributed by Susan L. Forsburg
Current Protocols in Molecular Biology (2003) 13.14.1-13.14.3
Copyright © 2003 by John Wiley & Sons, Inc.
13.14.1
Supplement 64
G1
S
M
G2 phase
Figure 13.14.1 The fission yeast cell cycle. Rod-shaped cells grow at the ends, maintaining
constant diameter. In exponentially growing cells, cytokinesis occurs after daughter nuclei have
already entered their next S phase.
Overview of
Schizosaccharomyces pombe
cerevisiae are available in S. pombe versions
that accommodate the distinct biology of the
fission yeast, and similar genetic strategies are
available for both systems (e.g., Forsburg,
2001). Fission yeast still lacks well developed
genomics resources, presumably because its
genome sequence was only completed in 2002.
Commercial microarrays or oligonucleotide
libraries are not widely available, and there is
as yet no standard collection of disruptions of
all open reading frames. However, other community-wide resources have been developed,
including libraries of temperature-sensitive
and insertion mutants at FYSSION at the University of Sussex, UK (http://pombe.biols.
susx.ac.uk), which may be freely screened. A
large expression project is underway at the
Sanger Centre and the data are freely searchable
(http://www.sanger.ac.uk/PostGenomics/S_pombe).
S. pombe cells are rod shaped and grow by
increasing length while maintaining a constant
diameter. The length of a cell is a sensitive
indicator of its position within the cell cycle.
They divide by medial fission, producing two
essentially identical daughter cells. The nuclear
cell cycle is divided into distinct G1 (10%), S
(10%), G2 (70%), and M (10%) phases. Following mitosis, the newly replicated nuclei enter the next cell cycle and undergo G1 and S
phase prior to completion of the previous cycle’s cytokinesis. This quirk means that a single
cell particle almost always has a 2C DNA
content, either because of the extended G2
phase, or the binucleate G1 or S phase cells (see
Fig. 13.14.1). Fission yeast is generally haploid
even in the wild. Following conjugation, newly
formed zygotes immediately enter meiosis and
sporulation to produce four spores in a linear
tetrad ascus. Diploids can be recovered in the
laboratory by selection for complementing
markers, but are unstable and prone to sporulate
even on minimal media. One of the major
differences in S. pombe and S. cerevisiae is in
handling mating and diploids (see UNIT 13.16).
Approximately 43% of genes have at least
one intron and multiple introns are common;
however, the introns are usually short, and most
are <100 bp in length (Wood et al., 2002). They
do not have the rigid splicing consensus observed in S. cerevisiae. TATA boxes and other
consensus sequences for transcription are more
proximal to the ATG than observed in budding
yeast (Russell, 1983). For this reason, it cannot
be assumed that fission yeast genes will express
appropriately in budding yeast, or vice versa.
Thus, for cross-complementation experiments
it is best to use a cDNA and a promoter native
to the host species. Plasmids and markers for
S. pombe are conceptually similar to those in S.
cerevisiae, with the exception of single-copy
centromere plasmids. Since the fission yeast
centromeres are so large, they cannot be accommodated on a typical episome. While some S.
cerevisiae plasmids and markers can be maintained in S. pombe, and were widely used in the
early days of fission yeast molecular biology,
they are not optimal and are prone to plasmid
loss or rearrangement. Plasmids containing S.
13.14.2
Supplement 64
Current Protocols in Molecular Biology
pombe–specific replication origins and markers are strongly recommended and a large variety now exist.
Most protocols described in the sections on
S. cerevisiae are broadly applicable to fission
yeast with only minor changes. The author’s
purpose here is to identify those common S.
pombe–specific methods that are required by a
new investigator but are substantially different
from similar protocols in budding yeast, including specific media (UNIT 13.15), crosses and diploids, tetrad analysis, and cell cycle synchronization (UNIT 13.16), and some types of transformation (UNIT 13.17). General handling of the
organism, microbial culture, and overall purification of nucleic acid and protein are similar
to budding yeast. Generally, an investigator
who has handled E. coli in the course of performing standard molecular biology manipulations will have no trouble working with yeast
of either species. Additional resources for
working with S. pombe can be found in
(Moreno et al., 1991; Alfa et al., 1993; Gould,
2003).
Mitchison, J.M. 1990. The fission yeast, Schizosaccharomyces pombe. BioEssays 12:189-191.
Moreno, S., Klar, A., and Nurse, P. 1991. Molecular
genetic analysis of the fission yeast Schizosaccharomyces pombe. Methods Enzymol. 194:795823.
Nurse, P. 2000. A Long Twentieth Century of the
Cell Cycle and Beyond. Cell 100:71-78.
Russell, P.R. 1983. Evolutionary divergence of the
messenger-RNA transcription initiation mechanism in yeast. Nature 301:167-169.
Wood, V., Gwilliam, R., Rajandream, M., Lyne, M.,
Lyne, R., Stewart, A., Sgouros, J., Peat, N.,
Hayles, J., Baker, S., et al. 2002. The genome
sequence of the eukaryote fission yeast Schizosaccharomyces pombe. Nature 415:871-880.
Yanagida, M. 2002. The model unicellular eukaryote, Schizosaccharomyces pombe. Genome
Biol. 3:2003.1-2003.4.
INTERNET RESOURCES
http://www.sanger.ac.uk/Projects/S_pombe
The Sanger Center S. pombe project.
http://www.genedb.org/genedb/pombe/index.jsp
The Sanger Center S. pombe GeneDB project.
LITERATURE CITED
http://www.sanger.ac.uk/PostGenomics/S_pombe
Alfa, C., Fantes, P., Hyams, J., McLeod, M., and
Warbrick, E. 1993. Experiments with Fission
Yeast. Cold Spring Harbor Laboratory Press,
Cold Spring Harbor, N.Y.
The Sanger Center expression analysis page.
Egel, R. 2000. Fission yeast on the brink of meiosis.
BioEssays 22.9:854-860.
Forsburg, S.L. 1999. The best yeast. Trends Genet.
15:340-344.
Forsburg, S.L. 2001. The art and design of genetic
screens: Yeast. Nat. Rev. Genet. 2:659-668.
Gould, K.L. (ed.) 2003. Methods for Schizosaccharomyces pombe. Methods Vol In Press.
http://www.pombe.net
The Forsburg Laboratory S. pombe pages.
http://pombe.biols.susx.ac.uk
The FYSSION homepage.
Contributed by Susan L. Forsburg
The Salk Institute for Biological Studies
La Jolla, California
Hedges, S.B. 2002. The origin and evolution of
model organisms. Nat. Rev. Genet. 3: 838-849.
Yeast
13.14.3
Current Protocols in Molecular Biology
Supplement 64
S. pombe Strain Maintenance and Media
UNIT 13.15
STRAIN MAINTENANCE AND GENERAL GUIDELINES
Growth conditions
The basic methods for yeast culture are the same for both S. pombe and S. cerevisiae (see
UNIT 13.2). The generation time of S. pombe is somewhat longer than that of budding yeast,
ranging from 2 to 5 hr depending upon the medium, strain, and temperature. The preferred
permissive temperature is 32°C, although cells grow well at 30°C. The maximum
permissive temperature is 36°C and the minimum is 17°C. Most temperature-sensitive
mutants are viable at 25°C.
Determining cell count
As an approximate guide, an OD600 of 1.0 equals ∼1.5 × 107 cells/ml, subject to the same
caveats described in UNIT 13.2; however, bear in mind that the OD is a measure of overall
cell mass, not cell number. A strain arrested by mutation in a cell cycle gene will continue
to increase in OD because the cells continue to elongate, even though the cell number is
not increasing. Under these conditions, a hemacytometer or Coulter counter should be
used to measure cell number.
Diploids
Diploid strains should not be maintained for prolonged periods, but constructed fresh
before use (see UNIT 13.16 on mating and mating type testing). The fission yeast diploid is
partially induced for meiosis, and prolonged growth leads to selection for mutants that
block the meiotic pathway. The exception is diploids that contain an otherwise lethal
disruption. It is common for diploids awakened from the freezer to have a high frequency
of sporulation-minus segregants; this can be examined by iodine staining, as described
below. Similarly, h+ strains which can revert to h90 should be tested to verify the mating
type. Yeast nomenclature is detailed in Table 13.15.1.
Table 13.15.1 Yeast Nomenclature
Species
Wild type gene
name
Recessive
mutant
Protein
Disruption
S. pombe
S. cerevisiae
yfg1+
YFG1
yfg1-1
yfg1-1
Yfg1, Yfg1p
Yfg1, Yfg1p
∆yfg1::ura4+; ∆yfg1-D1; yfg1∆ or ∆yfg1
∆yfg1::URA3; yfg1∆ or ∆yfg1
PRESERVING STRAINS BY FREEZING
Strains are typically stored at −70°C in glycerol. Stocks should be cultured to stationary
phase (OD600 = 1.5) in YES medium (see recipe), then mixed 1:1 either with sterile yellow
freezing mix (50% v/v glycerol in YES) or 50% (v/v) sterile glycerol and immediately
frozen. For strains that must be maintained under selection (e.g., to maintain a plasmid),
the culture should be grown in EMM (see recipe) and frozen by addition of an equal
volume of 50% (v/v) glycerol. Phenotypes of strains reawakened from the freezer should
be verified by replica plating.
BASIC
PROTOCOL
NOTE: Rather than sterile velvets, the authors recommend using two circles of Whatman
#1 filter paper, which are sterile straight out of the box.
Yeast
Contributed by Susan L. Forsburg
Current Protocols in Molecular Biology (2003) 13.15.1-13.15.5
Copyright © 2003 by John Wiley & Sons, Inc.
13.15.1
Supplement 64
MEDIA
Fission yeast can be grown on synthetic minimal or rich media (see recipes), depending
upon the needs of the experiment. Ideally, investigators should use media optimized for
the organism, although S. pombe can be maintained on budding yeast media if necessary.
Rich media made from yeast extract (YE) is usually supplemented to promote maximum
growth and prevent adenine limitation. There are several variations of minimal media,
although EMM (see recipe) is the most efficient at suppressing mating or sporulation.
There are multiple choices for mating media, including malt extract or SPAS (see recipe).
Most fission yeast investigators do not use “drop-out mix,” but add the individual
supplements required for their strains; however, this is a result of habit rather than a
requirement. Commercially available media mixes are also available from Qbiogene.
The same general rules apply for fission yeast media as for budding yeast media (UNIT
13.1), including sources for compounds, autoclaving, and general procedures. Media
should not be autoclaved for more than 20 min to prevent caramelization of the glucose.
Rich Medium
YES
Per liter:
5 g yeast extract
30 g glucose
225 mg adenine-HCl
225 mg L-histidine
225 mg L-leucine
225 mg uracil
225 mg L-lysine-HCl
20 g Difco Bacto Agar
Final concentration:
0.5% yeast extract
3% glucose
1.31 mM adenine
1.45 mM L-histidine
1.71 mM L-leucine
2.01 mM uracil
1.23 mM L-lysine
2% agar (solid medium only)
YES, or YE + supplements, is preferred for general growth when selection is not required;
however, when selecting for the kan-MX marker using G418/geneticin resistance (Sigma
G-5013), YES is the preferred media. The supplements are added to ensure maximum
growth. If the supplements are not added, normal YE is functionally a low-adenine media.
Note that YE or YES contains thiamine, so it represses expression from the commonly used
nmt1 (no message in thiamine) promoter.
Minimal Medium
EMM
Per liter:
3 g potassium hydrogen phthalate
2.2 g dibasic sodium phosphate
5 g ammonium chloride
20 g glucose
20 ml 50× salt stock (see recipe)
1 ml 1000× vitamin stock (see recipe)
0.1 ml 10,000× mineral stock (see recipe)
S. pombe Strain
Maintenance and
Media
Final concentration:
14.7 mM C8H5KO4
15.5 mM Na2HPO4
93.5 mM NH4Cl
2% glucose
1× salt
1× vitamins
1× minerals
When using Edinburgh minimal medium, required supplements for auxotrophies (e.g.,
adenine, uracil) are added immediately before use to a final concentration of 225 mg/liter
as required. These can be maintained as sterile stock solutions at 7.5 mg/ml in water (3.75
mg/ml for uracil). Low adenine media, which allows the development of a red color in Ade−
strains, reduces the amount of adenine to 7.5 mg/liter. Some protocols use 10 or even 30
mg/liter adenine, and still see satisfactory red color develop.
13.15.2
Supplement 64
Current Protocols in Molecular Biology
The nmt1 promoter, which is the most commonly used promoter for heterologous gene
expression (Maundrell, 1990, 1993), is repressed by the addition of 15 ìM thiamine (15
ìM or 5 ìg/ml; from filter sterilized stock at 10 mg/ml) immediately prior to use. The levels
of expression can also be titrated using intermediate amounts of thiamine (0.05 ìM or
0.016 ìg/ml; Javerzat et al., 1996). SD media, used for budding yeast, contains thiamine
and therefore cannot be used for fission yeast when nmt1 expression is desired.
Variations of EMM media are used for particular purposes. For example, EMM lacking
nitrogen (i.e., which leaves out the ammonium chloride; EMM −N) is used to arrest cells
in the G1 phase of the cell cycle at 25°C. EMM in which the ammonium chloride is replaced
by 1 g/liter L-glutamate is a low nitrogen medium that promotes sexual differentiation in
lieu of mating media discussed below. Under conditions where slow growth is desired to
allow full expression of a particular phenotype, ammonium chloride is often replaced with
a poor nitrogen source, for example 3.8 g/liter glutamate (Ekwall et al., 1996) or 1.2 g/liter
proline (Rhind and Russell, 1998). EMM with 0.5% glucose instead of 2% reportedly
improves some transformation procedures (Okazaki et al., 1990). EMM that lacks both
nitrogen and glucose (EMM −N −G) can be used to arrest cells in the G2 phase of the cell
cycle and is also a suitable buffer to maintain spores following random spore analysis (UNIT
13.16). EMM + sorbitol plates are used for plating protoplasts. Add sorbitol to a final
concentration of 1.2 M in standard EMM prior to autoclaving.
Stock Solutions
Prepare all stock solutions in water. Filter sterilize and store indefinitely at 4°C.
50× salt stock
Per liter:
52.5 g magnesium chloride hexahydrate
0.735 g calcium chloride dihydrate
50 g potassium chloride
2 g disodium sulfate
Final concentration:
0.26 M MgCl2⋅6 H2O
5.0 mM CaCl2⋅2 H2O
0.67 M KCl
4.1 mM Na2SO4
1000× vitamin stock
Per liter:
1 g pantothenic acid
10 g nicotinic acid
10 g inositol
10 mg biotin
Final concentration:
81.2 mM pantothenic acid
81.2 mM nicotinic acid
4.20 mM inositol
40.9 µM biotin
10,000× mineral stock
Per liter:
5 g boric acid
4 g magnesium sulfate
4 g zinc sulfate heptahydrate
2 g ferric chloride hexahydrate
0.4 g molybdic acid
1 g potassium iodide
0.4 g cupric sulfate pentahydrate
10 g citric acid
Final concentration:
80.9 mM boric acid
33.2 mM MnSO4
13.9 mM ZnSO4⋅7 H2O
7.40 mM FeCl2⋅6 H2O
0.32 mM molybdic acid
6.02 mM KI
1.60 mM CuSO4⋅5 H2O
47.6 mM citric acid
Yeast
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Mating Media
ME
Per liter:
30 g Bacto-malt extract (ME)
30 g glucose
225 mg adenine-HCl
225 mg L-histidine
225 mg L-leucine
225 mg uracil
20 g Difco Bacto agar
Adjust to pH 5.5 with NaOH
Final concentration:
3% (w/v) malt extract
3% glucose
1.31 mM adenine
1.45 mM L-histidine
1.71 mM L-leucine
2.01 mM uracil
2% agar (for solid medium only)
ME is the most common mating media, but may show batch-to-batch variation. More
defined media can be used instead, such as SPAS, EMM −N, or EMM with 1 g/liter
L-glutamate instead of NH4Cl.
SPAS
Per liter:
10 g glucose
1 g potassium phosphate monobasic
1 ml 1000× vitamin stock
45 mg adenine-HCl
45 mg histidine
45 mg leucine
45 mg uracil
45 mg lysine-HCl
30 g Difco Bacto agar
Final concentration:
1% (w/v) glucose
7.3 mM KH2PO4
1× vitamins
0.26 mM adenine (1/5 normal)
0.29 mM histidine (1/5 normal)
0.34 mM leucine (1/5 normal)
0.40 mM uracil (1/5 normal)
0.25 mM lysine (1/5 normal)
3% agar (for solid medium only)
Phloxin B
This vital stain is commonly used to determine ploidy and temperature sensitivity of S.
pombe. It colors solid media vivid pink and the colonies will range in color according to
the health of the cells. However, because it is mildly toxic, it should only be used for
testing and not for long-term storage. Cells on phloxin B media die rapidly if the plates
are refrigerated. Wild-type haploids form pale pink colonies on phloxin B. Diploid
colonies are slightly darker. Dying cells form darker colonies or patches, and lethal
mutants are often dark red. The color is not apparent at the single cell level (any dead cells
will stain vivid pink), but relies on macroscopic examination of colony color.
Phloxin B (Magdala Red; Sigma) is prepared at a final concentration of 5 mg/liter from
a stock of 10 g/liter in water. It is then filter sterilized and kept indefinitely at room
temperature. Because it is light sensitive, it should be stored in the dark (e.g., a foil-covered
tube). Plates should be stored at room temperature, in the dark, for no more than a few
weeks. If the plates start to dull or look brownish, they are past their usefulness.
LITERATURE CITED
Ekwall, K., Nimmo, E.R., Javerzat, J.P., Borgstrom, B., Egel, R., Cranston, G., and Allshire, R. 1996.
Mutations in the fission yeast silencing factors clr4+ and rik1+ disrupt the localisation of the chromo
domain protein Swi6p and impair centromere function. J. Cell. Sci. 109:2637-2648.
Javerzat, J.P., Cranston, G., and Allshire, R.C. 1996. Fission yeast genes which disrupt mitotic chromosome
segregation when overexpressed. Nucl. Acids Res. 24:4676-4683.
S. pombe Strain
Maintenance and
Media
Maundrell, K. 1990. nmt1 of fission yeast: A highly transcribed gene completely repressed by thiamine. J
Biol. Chem. 265:10857-10864.
13.15.4
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Current Protocols in Molecular Biology
Maundrell, K. 1993. Thiamine-repressible expression vectors prep and prip for fission yeast. Gene 123:127130.
Okazaki, K., Okazaki, N., Kume, K., Jinno, S., Tanaka, K., and Okayama, H. 1990. High-frequency
transformation method and library transducing vectors for cloning mammalian cDNAs by trans-complementation of Schizosaccharomyces pombe. Nucl. Acids Res. 18:6485-6489.
Rhind, N. and Russell, P. 1998. The Schizosaccharomyces pombe S-phase checkpoint differentiates between
different types of DNA damage. Genetics 149:1729-1737.
Contributed by Susan L. Forsburg
Molecular and Cell Biology Laboratory
The Salk Institute for Biological Studies
La Jolla, California
Yeast
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Supplement 64
Growth and Manipulation of S. pombe
UNIT 13.16
Much of the difference in working with S. pombe compared to S. cerevisiae comes from
two issues. First, S. pombe has a G2-oriented cell cycle, spending 70% of its lifecycle in
G2—almost completely opposite to the case of S. cerevisiae. Second, a wild-type cell is
usually haploid, because S. pombe cells mate and sporulate only in response to nutrient
limitation in a continuous pathway from conjugation through meiosis. This contrasts with
wild-type S. cerevisiae which will mate whenever an appropriate partner is present, but
only sporulates under nutrient limitation. Thus, working with fission yeast diploids in the
laboratory requires specific genetic tricks and planning ahead. Moreover, mating type
pheromones cannot be used to arrest the cell cycle easily in S. pombe, so other methods
must be employed. This unit addresses these distinct areas of working with crosses,
tetrads, and diploids, as well as analyzing the cell cycle and the products of genetic
analysis.
CROSSES, TETRADS, AND DIPLOIDS
Fission yeast has two mating types, h+ and h−, which are alleles of the mat1 locus (mat1-M
for h−, mat1-P for h+). Fully wild-type cells switch between h+ and h− mating types; these
are called h90 (because 90% of cells can typically switch). In a culture of h90 cells,
approximately half of the cells with be functionally h+ and half h− because of switching.
Laboratory investigators typically use strains with rearrangements of the silent mating
loci to provide a stable mating type and prevent switching. These heterothallic strains will
only mate if a partner of the opposite mating type is provided to them. Since the silent
loci are closely linked to the expressed mat1 locus, it is convenient (but not quite accurate)
to think of mating type in S. pombe as having three alleles: h90, h+, and h−. The common
h− is stable, but most laboratory h+ strains revert to h90 at a low frequency (∼10−3; for
review, see Klar, 1992).
The S. pombe lifestyle is predominantly haploid, and in contrast to budding yeast, fission
yeast haploids must be starved of nitrogen in order to undergo mating. Once they mate,
they normally proceed immediately through sporulation, generating a curved, zygotic
ascus (Fig. 13.16.1A). Diploids can be recovered in the laboratory by interrupting this
process, and can be maintained vegetatively; however, they are prone to sporulate, and
when they do, they form a linear, azygotic ascus that is easily distinguished from the
zygotic ascus (Fig. 13.16.1A).
In this unit, mating type testing is described (see Basic Protocol 1), crossing strains for
tetrad or random spore analysis is covered (see Basic Protocol 2), and instructions for
working with diploids are given (see Basic Protocol 3).
TETRAD DISSECTION AND RANDOM SPORE ANALYSIS
Both tetrad dissection and random spore analysis (RSA) are used to construct new strains
and for analysis of meiotic products. Because mating and meiosis are linked in S. pombe,
it is generally not necessary to isolate a diploid prior to characterizing offspring. Instead,
the haploid parents are crossed on mating plates and allowed to proceed all the way
through sporulation. The only exceptions are generally when very efficient sporulation is
required, or when one parent is homothallic (h90), and would otherwise cross with itself.
The logic of tetrad dissection is exactly the same for S. pombe as S. cerevisiae, and the
manipulations are similar. RSA in fission yeast is much simpler than for budding yeast
Yeast
Contributed by Susan L. Forsburg
Current Protocols in Molecular Biology (2003) 13.16.1-13.16.17
Copyright © 2003 by John Wiley & Sons, Inc.
13.16.1
Supplement 64
because vegetative cells are easily killed by the enzyme glusulase (snail gut enzyme), and
the spores do not stick to one another, so they are readily dispersed. RSA provides certain
advantages because it allows the analysis of a large population of offspring with relatively
little labor, and is very commonly used. On the other hand, tetrad dissection uniquely
allows the analysis of the products of an individual meiosis, and the unambiguous
identification of double mutants. S. pombe tetrads are generally somewhat linear in shape,
and therefore ordered, but it is seldom possible to dissect the spores in such as way as to
maintain the order.
How does one choose which is most useful? For general strain construction in which the
desired double mutant can be unambiguously identified, RSA is the method of choice. It
is particularly useful if the two mutations are linked (e.g., leu1 and his7), because a large
number of offspring can be analyzed to identify a rare event. RSA can be used for linkage
analysis in genetic screens, since large numbers of crosses can be processed easily, and
it is commonly used to determine recombination frequency in large mapping studies. The
only caveats are that all the genotypes must be equally viable and the desired recombinant
must be unambiguously identified. In cases where there is some doubt about the genotype
of a putative double mutant, the candidates can be backcrossed to each parent to verify
linkage. Most common strain construction requirements are satisfied by RSA.
There are two special cases where RSA is particularly helpful. The first is in recovery of
spores containing plasmids. Because fission yeast plasmids cannot encompass the very
large fission yeast centromere (UNIT 13.14), plasmids are meiotically unstable and usually
only 10% of spores will contain a plasmid marker. However, using RSA, even this low
fraction is sufficient to isolate a plasmid-containing spore or to draw conclusions about
plasmid complementation of deletion alleles in cases where plasmid shuffle is not
possible.
The second special case is the analysis of lethal mutations in the population during a spore
germination experiment. Diploid strains heterozygous for a marked allele are sporulated
in batch culture of liquid mating medium to generate large populations of spores.
Following digestion with the enzyme, the spores can be inoculated into selective medium
so that only cells carrying a desired marker are able to germinate. The rest of the spores
are inert so that the phenotype of the germinated mutants may be unambiguously
determined.
Tetrad analysis provides an enormous amount of information even in ten tetrads. Linkage
trends are usually visible in just a few tetrads (but meaningful numbers for weak linkage
may take many more). Most importantly, tetrad analysis allows the unambiguous identification of double mutants and is essential if the phenotype of a double mutant is unknown
or potentially lethal. Consider a cross between two viable mutants, ∆yfg1::ura4+ and
∆yfg1::ura4+. Since both are marked with ura4+ and are otherwise indistinguishable, the
investigator relies upon isolation of the nonparental ditype tetrad (2 ura4+:2 ura4− spores)
to identify unambiguously the double mutant. In this tetrad, the two ura4+ spores must
both be double mutants and their phenotypes and viability can be determined.
Wild-type fission yeast are efficient at germinating on YES plates. Germination is
modestly reduced on EMM plates, so unless selection for specific markers is required,
YES plates are preferred both for RSA and tetrad dissection.
Growth and
Manipulation of
S. pombe
In this unit, tetrad analysis is described (see Basic Protocol 4). Additional protocols detail
RSA (see Alternate Protocol 1), nonsporulating diploids (see Alternate Protocol 2), and
endoreduplication (see Alternate Protocol 3).
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Cell Cycle Synchronization
In budding yeast, the mating pheromone α factor is sufficient to arrest cells in G1 phase.
In wild-type fission yeast, mating pheromone has much weaker effects, and the primary
G1 arrest signal is provided by nitrogen starvation. Thus, cells can be arrested in G1 by
starving for nitrogen and then releasing by refeeding with complete medium. This leads
to a reasonably synchronous cell cycle following a several hour delay (lag phase).
Alternative methods for cell synchronization fall into two classes: those that separate
small G2 cells on the basis of size by some form of centrifugation and those that rely on
cell cycle block and release by starvation, drugs, or mutants. The latter are generally
simpler to perform but, since they rely on a perturbation of the cell cycle, they may lead
to confusing results. Any meaningful cell cycle variation in phenotype, protein, or RNA
levels should be verified using cycling cells unperturbed by arrest.
In these protocols the authors describe methods used for physiological analysis, such as
cell cycle dynamics, using small cultures. They can be adjusted for preparative studies of
RNA or protein by increasing the culture volume and harvesting larger aliquots at each
time point.
To determine cell cycle synchronization during nitrogen starvation, a time course is
monitored by sampling cells at each time point and fixing the aliquot for microscopy (see
Basic Protocol 5 and Support Protocol). Similar monitoring is used for other synchrony
methods, including cell cycle block and release (see Alternate Protocol 4). Size selection
of cells via lactose gradients is also described (see Basic Protocol 6). Use of centrifugal
elutriation requires a specialized centrifuge and rotor that is not found in most labs;
however, should such an instrument be available, a good description of the procedure is
found in Alfa et al. (1993).
MATING TYPE TESTING
Unlike S. cerevisiae, mating type testing in S. pombe relies upon completion of sporulation
rather than isolation of complementing diploids. This reflects the relative instability of
diploids in the laboratory. Thus, the only markers that are relevant in the strains to be
tested are their mating types. When fission yeast cells complete meiosis and sporulation,
they produce starch that can be stained easily by exposure to iodine vapor; however, iodine
vapor kills the cells, so this method cannot be used if further analysis of the meiotic
products is required.
BASIC
PROTOCOL 1
Materials
Known h+ and h− tester strains
Rich medium plates (e.g., YES plates; UNIT 13.15)
Characterized (positive control) and uncharacterized strains
Mating plates (ME, SPAS, or EMM −N; UNIT 13.15)
Iodine crystals (Sigma)
25°C incubator
Empty petri dish
Additional equipment and reagents for replica plating (UNITS 1.3 & 13.2)
CAUTION: Iodine is toxic and the vapors from the crystals or the exposed tester plate
should not be inhaled.
1. Patch known h+ and h− tester strains vertically on rich medium plates, and the
uncharacterized strain on a separate plate horizontally (Fig. 13.16.1). Include a
characterized strain as a positive control. Incubate at 32° (wild type) or 25°C (ts
strains) until both patches are grown up (1 to 2 days).
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A
B
zygotes
(from conjugated cells)
tester strains
unknowns
h+ h-
replica plate to
ME or SPAS
zygotic ascus
(from conjugated cells)
iodine stain
azygotic ascus
(from stable diploid)
hh+
hh90
h+
h+ h-
Figure 13.16.1 Fission yeast mating. (A) Zygotes and asci as seen under the microscope. (B)
Mating type testing on plates.
2. Replica plate onto mating plates so that they intersect (UNITS 1.3 & 13.2).
Cells mate most efficiently when they are replica plated from freshly grown streaks on rich
medium.
3. Incubate 3 to 4 days at 25° to 29°C.
Mating and sporulation are inherently temperature sensitive and should be carried out at
25°C (maximum of 29°C).
4. Prepare an iodine plate by scattering iodine crystals on the bottom or lid of an empty
petri dish in a fume hood. Remove the lid of the tester plate and invert over the plate
of crystals. Incubate this way until the positive control becomes dark brown where
the streak intersects the tester of the opposite mating type (5 to 10 min). Do not allow
the crystals to touch the agar or cells.
The plate with crystals can be stored in a sealed plastic container in the fume hood. The
crystals will continue to sublimate and will need to be refreshed regularly. If the tester plate
is left too long, the color can be reduced by removing the plate from the crystals and letting
it fade in the hood.
5. Determine the mating type by identifying the junctions in which a strain of known
mating type generated a dark patch with the strain being tested (see Fig. 13.16.1B).
h90strains will mate with all testers and themselves, generating a continuously dark
streak. The dark color does not last, so the results should be recorded immediately.
Growth and
Manipulation of
S. pombe
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CROSSING STRAINS FOR TETRAD OR RANDOM SPORE ANALYSIS
Construction of new strains or analysis of meiotic products requires that the parent strains
cross efficiently under conditions where the spores are viable. The previous protocol (see
Basic Protocol 1) is adequate for assigning mating type, but mating via replica plates is
relatively inefficient, and the iodine staining will kill the cells. This protocol provides
more efficient mating by improving the mixing of the parent cells, and does not employ
iodine. Successful mating can be determined microscopically by determining the formation of zygotes and asci (Fig. 13.16.1A).
BASIC
PROTOCOL 2
Materials
Strains to be crossed (i.e., strains 1 and 2) growing robustly on YES plates (EMM
if selection is required; UNIT 13.15)
Mating plates (ME or SPAS plates; UNIT 13.15)
H2O, sterile
Sterile toothpicks
25° to 29°C incubator
For single matings
1a. Using a sterile toothpick, take a visible glob of cells from strain 1 and make a small
patch on a mating plate (∼5 mm in diameter). Using a second sterile toothpick, take
an equivalent amount of strain 2 and add to the previous patch.
2a. Add 5 µl sterile water and mix strains gently on the surface of the agar with a sterile
toothpick.
For multiple matings using the same strains
1b. Pick a generous amount into 100 µl sterile water.
2b. Pipet 5 µl of each onto a patch and use a sterile toothpick to mix each cross gently.
Strains growing in liquid medium can be mated, but they must be washed free of the medium
or the nitrogen in the medium will suppress mating.
3. Incubate at 25° to 29°C for 2 to 4 days (2 to 3 for tetrads, 3 to 4 for other assays).
Monitor formation of asci microscopically.
Note that iodine staining (see Basic Protocol 1) is lethal and cannot be used if viable asci
are required for tetrad dissection, RSA, or spore germination.
Mating and sporulation are inherently temperature sensitive and should be carried out at
25°C (maximum of 29°C). Sterile strains can be forcibly mated using protoplast fusion, a
variation of the protoplast protocol used for transformation (see section on transformation
in UNIT 13.17). Mating and sporulation can also occur in liquid culture. Equal numbers of
both mating types, or previously isolated diploid cells, are washed free of normal growth
medium and then inoculated into liquid mating medium. Cultures are incubated with gentle
shaking at 25°C and monitored microscopically for the formation of zygotes and/or spores.
Generally this is accompanied by substantial flocculation.
WORKING WITH DIPLOIDS
Although fission yeast diploids are unstable, they can be recovered by interrupting the
mating process. Diploid strains must be selected by complementation of nutritional
markers; however, after they are identified, they should be maintained on YES to suppress
sporulation. Diploids are important for many assays and take only a little advance
planning to be useful in the laboratory.
BASIC
PROTOCOL 3
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A particularly convenient pair of markers for diploid construction is provided by two
complementing ade6 alleles, ade6-M210 and ade6-M216. When present in the same cell,
these alleles complement intragenically, leading to an Ade+ phenotype. This also provides
a color screen: the ade6+ gene is the ortholog of budding yeast ADE2, and mutants cause
the accumulation of a pink or red color in the cell in the presence of low adenine (see UNIT
+
13.15). Ade strains are white. The two ade6 alleles can be distinguished by their degree
of color: ade6-M210 is usually darker pink than ade6-M216. Neither is as dark as a null
allele; however, their color can be affected by the strain background, so the marker is
frequently misassigned. Assignment can be verified by attempting to make a diploid
against a strain with a known ade6 allele.
If the appropriate ade6 markers are unavailable, a diploid can be constructed by complementation of any two auxotrophic markers. In this case, the investigator must use care to
verify that the strain recovered is diploid, and not a recombinant haploid spore.
Materials
Strains to be crossed (complementing mating types and ade6 markers) growing
robustly on YES plates (EMM if required for selection; UNIT 13.15)
EMM plates lacking adenine (UNIT 13.15)
YES plates containing phloxin B (UNIT 13.15)
YES plates (UNIT 13.15)
Mating plates (ME or SPAS; UNIT 13.15)
Sterile toothpicks
25° or 32°C incubator
Additional reagents and equipment for mating and testing the mating type of S.
pombe (see Basic Protocol 1)
1. Follow the steps for mating S. pombe (see Basic Protocol 1, steps 1 to 3), using strains
with complementing mating types and ade6 markers growing robustly on YES (or
EMM) plates.
2. At 6 hr after mating and again at 12 hr, take a sterile toothpick and pick a swatch of
cells. Streak to EMM medium lacking adenine (but containing any other required
supplements).
3. Incubate at 32°C (25°C for temperature-sensitive diploids).
Incubation at 36°C, if the diploids are not temperature sensitive, will suppress sporulation.
4. Once clearly white colonies are visible (1 to 2 days), pick and streak several on YES
plates containing phloxin B. Incubate 2 to 4 days at 32°C (25°C for temperature-sensitive mutants).
The cells in the middle of a colony on EMM may have sporulated; these will form pale pink
haploid colonies when streaked on YES plates containing phloxin B. The darker pink
colonies are still diploids. It is useful to streak a known haploid on YES containing phloxin
B plates to verify its color. In addition, diploids are longer and wider than haploids when
viewed under the microscope.
5. Pick several independent colonies and streak to YES plates. Patch the same colony
in an ∼5- to 10-mm spot on mating plates (ME or SPAS) and incubate 1 to 2 days at
25°C. Expose the mating plates to iodine vapor as described in mating type testing
(see Basic Protocol 1) to verify clones are sporulation competent.
Diploids are stable for 1 to 2 weeks at 4°C if maintained on YES.
Growth and
Manipulation of
S. pombe
Prolonged maintenance selects for sporulation deficient mutants; therefore, diploids
should be constructed fresh whenever possible.
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TETRAD ANALYSIS
This method is almost identical to that in budding yeast and the apparatus is the same (see
UNIT 13.2). The significant difference is that fission yeast asci do not require prior digestion
with glusulase: they fall apart by themselves. Thus, the investigator must identify a mating
plate with ripe asci before they have started to disintegrate, and manipulate the intact asci
out to the position as shown in Figure 13.2.1. The plate is incubated for a few hours at
room temperature (or overnight at 17°C), during which time the ascus breaks down, and
the investigator returns to the plate and completes the dissection of the now freed tetrads.
This must be completed before the freed spores enter their first cell division. The
disintegration of asci is retarded at low temperatures and accelerated at high temperatures.
BASIC
PROTOCOL 4
In addition, fission yeast spores are not particularly sticky. Thus, a very high quality needle
with a completely flat plane is essential, since the spores will not stick to the needle
without surface tension. In addition to pulling needles oneself, precut fiber optic needles
that fit standard tetrad microscopes are also available (e.g., Singer Instruments;
http://www.singerinstruments.co.uk).
Materials
Mating/sporulation plate grown for 2 to 3 (haploid) or 1 to 2 (diploid) days
YES plates (UNIT 13.15)
Sterile toothpicks
Tetrad dissecting microscope
36°C incubator (optional)
17°C incubator (optional)
25° or 32°C incubator
Additional reagents and equipment for preparation and dissection of tetrads (UNIT
13.2)
NOTE: Azygotic asci from previously isolated diploids form more quickly than zygotic
asci from haploids, which must mate first.
1. Under a microscope, examine a mating/sporulation plate which has been grown for
2 to 3 (zygotic) or 1 to 2 (azygotic) days at 25° to 29°C to identify ripe asci, which
are intact but have clearly distinguished spores. If necessary, spread cells in the mating
patch along the agar with a sterile toothpick to make them more apparent.
The mating/sporulation plate can be left in the refrigerator for a day or two to retard ascus
breakdown if the investigator cannot immediately dissect the tetrads.
2. Transfer and lay cells across the YES plate upon which dissection will be performed
using a sterile toothpick.
3. Upon identification of a ripe intact ascus, use a dissecting needle to manipulate the
ascus to a fixed position (a of Fig. 13.2.1) on the plate (see protocol for preparation
and dissection of tetrads, UNIT 13.2). Place each additional ascus 0.5 cm to one side of
the preceding tetrad.
4a. For tetrads identified in the morning: Incubate the plate a few hours at warmer
temperatures (i.e., 32° to 36°C) to stimulate ascus breakdown. If the strain is not
temperature sensitive, an hour or two at 36°C can noticeably accelerate the process.
4b. For tetrads identified at the end of the day: Incubate the plate overnight at room
temperature or 17°C.
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5. Upon returning the plate to the micromanipulator, identify tetrads that have popped
by the four large, round, free spores and sometimes the faint skin of an ascus. Move
them down the plate to the b, c, and d positions (Fig. 13.2.1). Repeat steps 3 and 4
throughout the day as needed.
6. Once the plate is completed, incubate it at the appropriate temperature (32°C for wild
type, 25°C for temperature-sensitive mutants).
Germination occurs within a few hours and it generally takes 2 to 5 days to observe
colonies, depending upon the temperature of incubation.
ALTERNATE
PROTOCOL 1
RANDOM SPORE ANALYSIS
Fission yeast spores do not adhere to one another outside of the ascus; rather, the ascus falls
apart over time and the remaining vegetative diploids in a culture are readily killed by
glusulase. Thus, random spore analysis is significantly easier in S. pombe than in budding
yeast. It is the method of choice for many strain constructions, as long as the recombinant
strains are viable and easily distinguished from the parents. When used in bulk liquid culture,
it allows spore germination experiments to analyze large populations of marked, lethal alleles.
Materials
Mating/sporulation plate
H2O, sterile
5% glusulase (DuPont NEN) in sterile water
PBS (APPENDIX 2; optional), or EMM −N −G (UNIT 13.15; optional)
YES plates (UNIT 13.15)
Sterile toothpick
Additional reagents and equipment for counting cells using a hemacytometer
(APPENDIX 3F)
1. Using a sterile toothpick, pick cells from a mating/sporulation plate into 100 µl sterile
water. Add 10 µl of 5% glusulase and incubate overnight at room temperature.
2. Using a hemacytometer (APPENDIX 3F), count the number of spores in 10 µl of a 1:10
dilution in sterile water. Verify that no complete asci remain.
Typically all the spores in the central 5 × 5 grid area of an improved Neubauer hemacytometer are counted, and the number multiplied by 100 to give the number of spores per
milliliter.
3. Dilute in sterile water, PBS, or EMM −N −G and plate 500 spores per plate (assuming
all spores are viable). If selecting directly for spores containing particular markers,
adjust this number accordingly. If selecting directly for spores containing a plasmid,
plate 1000 to 10,000 spores per plate.
Do not use medium containing glucose for the dilutions, because it will induce germination.
The dilutions can be stored at 4°C for several days in case replating is required (e.g., if the
plates were plated too densely). Alternatively, spores in the original glusulase suspension
can be washed free of the enzyme in EMM −N −G and stored at 4°C for many days in case
further plating is required.
Colonies will appear in 2 to 5 days, depending on the temperature, and plates can be replica
plated to assign markers.
Growth and
Manipulation of
S. pombe
The same procedure can be employed at a larger scale on cultures of sporulated cells in
liquid medium, for a spore germination experiment. Following digestion (generally overnight or longer), the spores are washed several times before inoculation into selective
medium. Only the fraction of the population containing the selected marker(s) will
germinate. Any phenotype associated with the marker(s) can thus unambiguously be
distinguished from the inert background of wild-type spores.
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NONSPORULATING DIPLOIDS
+
−
Given the instability of normal h /h diploids, it can be useful to work with nonsporulating
diploids. Typically, these are either homozygous at the mating locus, or contain a mutation
in the h+information that prevents sporulation (Egel and Egel-Mitani, 1974; Willer et al.,
1995). These can sporulate by providing the missing mating information or by inducing
meiosis independent of mating type signaling by inactivating pat1 kinase. They can also
be induced to haploidize using drugs promoting chromosome loss (e.g., Bodi et al., 1991).
These strains have specialized uses including mapping genetic loci to chromosomes,
assessing the effect of ploidy on mutant phenotypes, or testing rates of chromosome loss.
ENDOREDUPLICATION
At a low frequency, wild-type fission yeast cells will occasionally skip mitosis and
produce a homozygous diploid, a process called endoreduplication. Because such diploids
are homozygous at the mating locus, they are incapable of sporulation; however, they are
still larger and darker on phloxin B. They can be isolated by screening cells plated on
YES containing phloxin B for darker pink colonies, which arise at a frequency of ∼10−3.
ALTERNATE
PROTOCOL 2
ALTERNATE
PROTOCOL 3
Materials
YES plates containing phloxin B (UNIT 13.15)
Liquid culture of strain of interest
1. Plate 500 to 1000 cells per plate on YES plates containing phloxin B.
2. Incubate at 25° or 32°C until colonies form and color develops (generally 2 to 5 days).
3. Pick darker pink colonies and restreak on YES plates containing phloxin B plates.
Verify diploid morphology by microscopy and ploidy by flow cytometry.
CELL-CYCLE SYNCHRONIZATION BY NITROGEN STARVATION
At low temperatures (25°C), S. pombe cells placed in EMM without nitrogen or supplements will arrest predominantly in G1. This is a temperature-sensitive effect, because at
higher temperatures, a larger fraction of the cells will arrest in G2 (Costello et al., 1986).
Auxotrophic markers such as ade6 may retard this arrest slightly, which can be accommodated by arresting the cells in EMM without nitrogen supplemented with 45 mg/liter
adenine (1/5 normal). Cells released from nitrogen by refeeding with regular EMM will
enter the cell cycle after ∼2 hr delay (the so-called “lag phase”). Nitrogen starvation cannot
be used to synchronize h90 or diploid cells, because absence of nitrogen triggers the sexual
differentiation process.
BASIC
PROTOCOL 5
Materials
Cells from the strain of interest
EMM and EMM −N (UNIT 13.15)
Complete medium (e.g., EMM with supplements, YES; UNIT 13.15)
25°C incubator
Incubator set at growth temperature (i.e., 32° to 36°C)
1. Grow cells from the strain of interest to mid-late exponential phase (OD = 0.6 to 1.0)
in EMM with nitrogen.
The amount of culture will depend upon the purpose of the experiment—i.e., whether it is
physiological analysis, in which case 50 ml will do, or synchrony to recover material for
protein or RNA analysis, in which case 300 to 500 ml are more appropriate.
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1 cell, septated
2 cells
2 cells
2 cells
Percent septated
1 cell
X
X
XX
X
X
X
X
X
XX X
XX
X
X
Time
Figure 13.16.2 Determining septation index for synchronous cultures.
2. Harvest by centrifuging 5 to 10 min at 4000 × g, 20°C. Wash twice with an equal
volume starvation medium.
3. Resuspend in an equal volume starvation medium (i.e., EMM −N) and incubate 12
hr at 25°C.
4. Add an equal volume prewarmed complete medium, preferably EMM containing any
required supplements. Incubate ∼2 hr at the appropriate growth temperature.
The temperature of the re-fed culture depends upon the purpose of the experiment. For a
normal wild-type strain, the cells should be returned to 32°C. For a temperature-sensitive
strain under restrictive conditions, the cells should be incubated at 36°C. It will take ∼2 hr
for cells released from nitrogen starvation in G1 to enter S phase.
YES can also be used to re-feed but, because it is a less defined medium subject to batch
variations, this is not preferred.
5. Every 20 to 30 min, take an aliquot of cells (∼5 µl) and place on a slide with a cover
slip. Examine under a phase microscope (dark-field illumination may help) and count
the percent septated cells (septation index), which corresponds to G1/S phase cells
(Fig. 13.16.2). Plot as a function of time.
The degree of synchrony is determined by the maximum peak of septation; generally 30%
for this method and up to 80% for the following method. This is ideally performed on live
cells to monitor the time course in real time. Because of the structure of the S. pombe cell
cycle (Fig. 13.14.1), the peak of septation in wild-type strains is roughly the same as cells
in S phase.
Aliquots may also be fixed in ethanol for FACS or microscopy and the septation index
counted from fixed cells if this is more convenient (see Support Protocol). The key to using
the septation index is to clearly distinguish cells that have begun division from those that
have not (Fig. 13.16.2). Consistency is essential so that in a given experiment, it is best if
a single investigator counts all the cultures. Preparative analysis requires larger aliquots;
cells can be harvested by centrifugation, the pellets frozen on dry ice, and stored for weeks
or longer at −70°C for subsequent analysis.
SUPPORT
PROTOCOL
Growth and
Manipulation of
S. pombe
ETHANOL FIXATION
Samples fixed in 70% ethanol may be stored indefinitely at 4°C. This is particularly useful
if FACS analysis or DAPI staining will also be performed, as the fixation is identical.
Materials
Exponentially growing culture
70% ethanol, 4°C
H2O, sterile
Mounting medium with stain (see recipe; optional)
Positively charged microscope slides
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1. Harvest 107 cells from an exponentially growing culture by centrifuging 5 min at
4000 × g, 20°C. Pour off the supernatant.
2. Vortex the tube while adding 1.0 ml cold 70% ethanol. Store at 4°C.
3. To rehydrate the cells, take 0.1 ml and add to 1 ml sterile water in a microcentrifuge
tube. Mix and microcentrifuge 5 min. Decant the supernatant and resuspend in 0.1
ml sterile water. If desired, process for septation index as described (see Basic
Protocol 5).
4. Optional: To visualize the nucleus and/or septa, pipet 5 µl cell suspension onto a
cleaned microscope slide. Allow to air dry or place on a heating plate at low
temperature until the liquid is just evaporated.
To ensure a monolayer of cells, pretreat the slides with poly-L-lysine (Sigma) or use a
positively charged slide.
5. Optional: Add 5 µl mounting medium with stain and immediately cover with a cover
slip. Examine using epifluorescence.
Samples can be stained with DAPI to visualize the nucleus, with Calcofluor to visualize
the septa, or processed for FACS analysis.
CELL CYCLE BLOCK AND RELEASE
Alternative methods of cell cycle synchronization rely upon reversible temperature-sensitive cell cycle mutants, with the cdc25-22 mutant being the most commonly used. It
arrests cells at the G2 phase of the cell cycle at 35.5°C (Russell and Nurse, 1986).
Following return to permissive temperature, cdc25 strains rapidly enter mitosis and
proceed through a synchronous cell cycle. The second cell cycle is significantly less
synchronous.
ALTERNATE
PROTOCOL 4
The key to temperature arrest is rapid increase and decrease of temperature. All water
baths should be prewarmed to the correct temperature. The volume of culture should be
low for the size of the flask, to ensure most efficient temperature exchange.
Materials
Temperature-sensitive cell cycle mutants (e.g., cdc25-22; ATCC# 90337)
25° and 36°C water baths
Thermometer cleaned with ethanol
1. Grow temperature-sensitive cell cycle mutants to early/mid-log phase (OD600 = 0.2
to 0.4) at permissive temperature. If necessary, remove an aliquot for an exponentially
growing cell time point, which can be processed for the same morphological or
molecular tests as the synchronized cells.
The volume taken will depend upon the assay: <1 ml for cell morphology or FACS, >10
ml for protein or RNA preparation.
2. Incubate 4 to 6 hr at 36°C.
A rapid temperature shift is important. Water baths promote the most efficient thermal
transfer. If a 36°C water bath is not available, swirl the flask in a 50°C bath or sink until
the culture approaches 36°C as described in step 3, then transfer to an air shaker.
3. Take an aliquot for a t = 0 time point. Release cells to 25°C by rapidly swirling the
flask in ice water and monitoring with a thermometer that has been cleaned with
ethanol. Place in 25°C water bath.
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4. To follow synchrony, sample time points every 20 to 30 min for septation index
(correlates with S phase). This should approach 60% to 80% for an effective
temperature-driven block and release.
Samples can be fixed for simple microscopy, or pellets can be harvested and frozen for RNA
or protein analysis. If complex manipulations are required, it is often easier to have two
people taking time points.
Because of their pronounced cell elongation, cdc25 mutants often give ambiguous FACS
profiles following release to permissive temperature.
BASIC
PROTOCOL 6
LACTOSE GRADIENT CENTRIFUGATION
Because fission yeast cells are very regular in size, size selection to isolate newborn G2
cells is an effective means to synchronize a population. Centrifugal elutriation is a frequent
method in fission yeast, but requires an expensive and highly specialized centrifuge rotor,
and working with more than one strain at a time is difficult. Lactose gradient centrifugation (e.g., Edwards and Carr, 1997) offers a convenient alternative that can be performed
in most laboratories on multiple strains. Both these methods rely on separating cells based
on cell size, so the strains must not show elongation or variation in cell morphology.
Mutants that have heterogeneous phenotypes or cause elongation are much more difficult
to synchronize using size selection methods.
Materials
Cells
70% ethanol
YES (UNIT 13.15) with 0%, 10%, and 40% lactose, 32°C
Gradient maker
20- and 50-ml conical tubes
Prepare culture
1. Grow a 125-ml culture overnight so that it is in mid- to late-exponential phase the
next day (OD600 = 0.5 to 0.8).
Prepare gradient
2. Sterilize a gradient maker by passing 70% ethanol through the entire apparatus, being
sure to loosen the screw so that both compartments empty into the tube. Place a tiny
stir bar in the compartment closest to the exit tube (Fig. 13.16.3), put the gradient
maker on a stir plate, and let the solutions drain down into a collection/waste beaker.
After the ethanol has run through the gradient maker, rinse with YES containing no
lactose.
3. Close the valve between the compartments. Load YES with 10% lactose in the
chamber away from the exit tube. Open the central valve briefly to clear any bubbles.
Load YES with 40% lactose in the compartment with the stir bar, proximal to the exit
tube. Again, clear bubbles. Turn on the stir plate so that the stir-bar in the 40%
compartment is mixing (Fig. 13.16.3).
4. Pour 5 ml YES with 40% lactose into a sterile 50-ml conical tube. Open both valves
of the gradient maker, allowing the gradient to flow down the sides of the conical
tube. Fill to 45 ml.
Growth and
Manipulation of
S. pombe
Once poured, the gradient is stable for an hour. Additional gradients can be poured if
required to allow processing of multiple strains.
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10%
40%
Figure 13.16.3 Assembling a lactose gradient.
Load culture onto gradient
5. Harvest cells by centrifuging at 3000 × g (∼5000 rpm), at 20°C until pelleted (5 to
10 min).
6. Resuspend cells in 0.5 ml YES with no lactose and pipet along the inside of the conical
tube just above the gradient solution.
7. Centrifuge the conical tube in a swinging bucket centrifuge 5 min at 228 × g (1000
rpm), room temperature.
8. Collect 5 ml from the top of the gradient in 1-ml aliquots using a pipettor. Examine
the cells in each aliquot using the microscope. Pool all aliquots.
When removing the aliquots, the tip of the pipet should barely touch the top of the gradient.
The goal is a uniform population of small G2 cells; no longer cells nor any septated cells
should be seen.
9. Spin down the aliquot of cells, wash in YES medium with 0% lactose, and resuspend
in 10 ml YES medium. Follow cell progression by septation and FACS.
A maximum septation index of ∼40% is expected.
10. Optional: If the above separation does not provide sufficient synchrony, or sufficient
cell number, repeat the procedure using a larger starting culture (500 ml).
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11. Optional: Further purify cells as follows:
a. Add 0.5 ml of 40% lactose in YES to the bottom of a 20-ml conical tube.
b. Add 10 ml YES/lactose from the gradient maker (step 3) on top of this initial
addition.
c. Layer the 5 ml harvested from the first gradient on the second, and centrifuge 3
min at 228 × g (1000 rpm), room temperature.
d. Recover six 0.5-ml aliquots, wash into fresh medium, and follow as above.
REAGENTS AND SOLUTIONS
Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see
APPENDIX 2; for suppliers, see APPENDIX 4.
Calcofluor white, 1 mg/ml
Prepare Calcofluor white (fluorescent brightener 28; Sigma) at a final concentration
of 1 mg/ml in 50 mM sodium citrate/100 mM sodium phosphate, pH 6.0 (APPENDIX
2). Store for months in the dark at 4°C.
DAPI stock
Dissolve 1 mg DAPI per milliliter DMSO. Store weeks to months in the dark at
−20°C. Discard if the solution turns dark brown.
Mounting medium stock
Dissolve the following in 5 ml PBS (APPENDIX 2):
50 mg n-propyl gallate (Sigma)
50 mg p-phenylenediamine (PPD; Sigma)
Add glycerol to 50 ml
Aliquot and store for months at −20°C in the dark
Discard if solution darkens
Mounting medium with stain
Staining can be performed with DAPI, Calcofluor white, or both, using the following
solutions.
DAPI alone
Dilute 1 µl of DAPI stock (i.e., 1 mg DAPI/1 ml DMSO; see recipe) per 1.5 ml
mounting medium stock (see recipe). Store 1 to 2 days at −20°C in the dark. Discard
if the solution turns brown.
Calcofluor white alone
Dilute 1 mg/ml Calcofluor white to 50 µg/ml in mounting medium stock (see recipe)
prior to use. Prepare fresh.
Calcofluor white and DAPI
Prepare a solution of mounting medium stock (see recipe) containing both stains as
described for the individual components. Prepare fresh. Note that the concentration
of Calcofluor white may need adjustment to prevent overcoming the DAPI signal.
COMMENTARY
Background information
Growth and
Manipulation of
S. pombe
Several important biological differences between S. pombe and S. cerevisiae are particularly worth noting. First, the fission yeast is
generally haploid, and can only be maintained
as a diploid with complementing markers or
other genetic tricks. Thus, linkage analysis is
often more efficient than complementation to
assess whether two mutations affect the same
gene. Second, because mating and meiosis are
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coupled, it is not necessary to isolate a stable
diploid prior to sporulating cells and analyzing
meiotic products: haploid strains are crossed
and allowed to proceed through meiosis in one
process. Third, the ease of random spore analysis in fission yeast makes isolation of even rare
meiotic products straightforward and, moreover, facilitates bulk sporulation and germination in liquid culture. The methods described
here are sufficient for the basic genetic manipulation of fission yeast in any laboratory.
Individual labs may use slightly different
protocols or medium, but in practical terms this
makes little difference in assessing mating type,
isolating diploids, or analyzing meiotic products by tetrads, RSA, or spore germination. For
additional background on the detailed biology
of mating type and meiosis, see Egel (2000),
Klar (1992), and Yamamoto (1996).
There is considerably more discussion about
the relative uses of different methods of cell
cycle synchronization (for general reviews of
cell cycle control, see MacNeill and Nurse,
1997; Moser and Russell, 2000). Methods that
arrest and then release the cell cycle, such as
nitrogen starvation (see Basic Protocol 5), cell
cycle block and release (see Alternate Protocol
4), or drug treatments (such as hydroxyurea,
which arrests early in S phase) allow very high
levels of synchronization—i.e., a septation index approaching 65% or more. However, in all
cases, there is substantial perturbation of the
cell cycle: just because some events are blocked
during the arrest, does not mean that all events
are blocked. For example, cdc25 mutants arrested in G2 phase continue to accumulate
cyclin to higher than normal levels (Moreno et
al., 1989). This is an important caveat that may
influence downstream interpretation of the results. Generally, the use of two independent
methods of synchronization is recommended
to verify that the results reflect normal cell cycle
variations, rather than an effect of arrest.
Not all mutants release as well as cdc25, but
well-behaved alternatives include cdc10-M17
(temperature sensitive, blocks at G1; Verkade
and O’Connell, 1998) and nda3-KM311 (cold
sensitive; blocks in mitosis; Umesono et al.,
1983). These may be employed similarly to
cdc25, although the temperatures are reversed
for nda3. For conditions where simple cell
cycle arrest is sufficient without release, cdc10,
cdc25, and nda3 mutants may be compared
with temperature-sensitive mutants affecting
other parts of the cell cycle, including cdc22
(ribonucleotide reductase; blocks in early S
phase; Fernandez Sarabia et al., 1993), cdc17
(DNA ligase, blocks late in S phase; Nasmyth,
1977), or nuc2 (APC component, arrests in
mitosis/G1; Hirano et al., 1988).
Size-selection methods of synchrony such
as lactose gradients (see Basic Protocol 6) or
elutriation (Alfa et al., 1993) do not overtly
perturb the cell cycle, but are less synchronous
than the arrest-release approaches. The maximum septation index is ∼35% to 40%, and the
peaks of septation are much broader. As described in UNIT 13.14, newly separated fission
yeast cells are in G2; therefore, methods that
preferentially isolate small cells will enrich the
G2 population, the basis for these procedures.
The cells may suffer some modest stress from
the low-speed centrifugation, but under conditions where temperature-sensitive mutants cannot be employed, or where normal cell cycle
responses are particularly important, these are
the methods of choice.
The reader is referred elsewhere for fundamentals of molecular analysis (Moreno et al.,
1991) and other methods in fission yeast (Alfa
et al., 1993; Gould, 2003).
Critical Parameters and
Troubleshooting
Mating
The mating type testing protocol (see Basic
Protocol 1) is often inefficient because it relies
upon relatively low numbers of cells transferred
by replica plating. Strains that grow slowly, that
have impaired starvation response, or that have
defects in mating may not generate sufficient
numbers of asci to be observed by iodine staining. Under these conditions, crossing strains for
tetrad or random spore analysis (see Basic Protocol 2) can be used and the plates subjected to
iodine staining, or monitored for zygote formation by microscopy. If the latter, a sterile toothpick is used to pull the cells out from the mating
patch and spread them out on the adjacent area
of the plate, or apply them to a microscope slide
in 5 µl sterile water. Zygotes and asci as in
Figure 13.16.1A can be observed under these
conditions. It is generally a good idea to set up
crosses for mating type testing, tetrads, or diploid isolation with a known pair of wild-type
strains to confirm that the medium and conditions are suitable for analysis.
Tetrads and RSA
It is a good idea to practice both tetrads and
RSA on well-behaved wild-type strains before
performing them on mutants of interest. In
addition, prior to replica plating, the micro-
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scope should be employed, especially on tetrad
plates where there are “missing” colonies, to
verify that there were actually spores manipulated in the gaps. This also provides evidence
of the phenotype prior to replica plating, when
single spores or dead cells can be lost.
Diploids
The most common reason for failure to recover a diploid is misassignment of the mating
type marker, or the ade6 marker. These can be
distinguished by observing the original mating
plate to see whether zygotes and asci formed.
If they do, then the likely culprit is the ade6
allele, which can be tested by setting up crosses
with a control strain with the opposite marker.
Stable diploids
Isolation of endoreduplicated strains to create stable homozygous diploids requires a
method to verify ploidy in addition to phloxin
B staining, because there is frequently a background of more darkly staining colonies that
are still haploid in the population. FACS is
unambiguous if a known haploid and diploid
are used as controls. Isolation of a haploid from
the stable homozygous diploid relies upon
chromosome loss. If fission yeast diploids lose
one chromosome, they will rapidly haploidize
and this can be accelerated with certain drugs
(e.g., Bodi et al., 1991).
An alternative method to create stable diploids relies upon crossing two strains with complementing markers which are able to mate but
not sporulate. The mat1-P mating type locus
produces two transcripts, one required for mating, and the second for meiosis (Egel and EgelMitani, 1974; Willer et al., 1995). A homothallic strain containing the mat2-102 allele lacks
the sporulation-specific P transcript. By itself
on mating medium it makes zygotes but no
spores. If mated as an h+ to a wild-type h− and
selected for complementing markers, the resulting diploid is unable to enter meiosis. In
contrast, the mat2-102 strain functions as a
normal h−, so if mated to a wild-type h+, the
resulting diploid is sporulation-competent. In
this way, the mat2-102 allele can be moved into
different backgrounds.
Growth and
Manipulation of
S. pombe
Synchronization
The success of synchronization procedures
is heavily influenced by the strain background.
Not all mutants will arrest appropriately in G1
phase when starved for nitrogen. Similarly,
many mutants cause a heterogeneous size phenotype that precludes use of size-selection
methods. These difficulties are only established
by trial and error. The experiment should be
performed on small-scale cultures, monitoring
septation index and FACS if appropriate, to see
if conditions are suitable before scaling up to
preparative levels. Often it helps to divide the
work between two people (one to harvest and
fix cells, one to count septa).
Anticipated Results
Crosses, diploids, and tetrads are the fundamental components of classical genetic analysis. The methods are straightforward and allow
determination of synthetic phenotypes or construction of useful strains. Synchronization
methods are important because of S. pombe’s
continuing importance as a model system for
cell cycle analysis.
Time Considerations
For cell growth experiments, the limiting
factor is growth rate of the individual strain.
This varies substantially depending upon the
genotype, medium, and temperature. Generation of recombinant strains takes about a week
from crossing the haploid parents, to replica
plating the germinated spores. Isolation and
purification of a diploid strain generally takes
about a week. Synchronous cultures generally
take all day to perform, not including time for
preparative analysis.
Literature Cited
Alfa, C., Fantes, P., Hyams, J., McLeod, M., and
Warbrick, E. 1993. Experiments With Fission
Yeast. Cold Spring Harbor Laboratory Press,
Cold Spring Harbor, N.Y.
Bodi, Z., Gysler-Junker, A., and Kohli, J. 1991. A
quantitative assay to measure chromosome stability in Schizosaccharomyces pombe. Mol. Gen.
Genet. 229:77-80.
Costello, G., Rodgers, L., and Beach, D. 1986. Fission yeast enters the stationary phase G0 state
from either mitotic G1 or G2. Curr. Genet.
11:119-125.
Edwards, R.J. and Carr, A.M. 1997. Analysis of
radiation-sensitive mutants of fission yeast.
Methods Enzymol. 283:471-493.
Egel, R. 2000. Fission yeast on the brink of meiosis.
BioEssays 22.9:854-860.
Egel, R. and Egel-Mitani, M. 1974. Premeiotic DNA
synthesis in fission yeast. Exp. Cell. Res. 88:127134.
Fernandez Sarabia, M.-J., McInerny, C., Harris, P.,
Gordon, C., and Fantes, P. 1993. The cell cycle
genes cdc22+ and suc22+ of the fission yeast
Schizosaccharomyces pombe encode the large
and small subunits of ribonucleotide reductase.
Mol. Gen. Genet. 238:241-251.
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Gould, K.L. (ed.) 2003. Methods for Schizosaccharomyces pombe. Methods Vol In Press.
Hirano, T., Hiraoka, Y., and Yanagida, M. 1988. A
temperature-sensitive mutation of the Schizosaccharomyces pombe gene nuc2+ that encodes a
nuclear scaffold-like protein blocks spindle
elongation in mitotic anaphase. J. Cell Biol.
106:1171-1183.
Klar, A.J.S. 1992. Molecular genetics of fission
yeast cell type: Mating type and mating type
interconversion. In The Molecular and Cellular
Biology of the Yeast Saccharomyces: Gene Expression (E. Jones, J. Pringle, and J. Broach,
eds.) pp. 745-777. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
MacNeill, S.A. and Nurse, P. 1997. Cell cycle control in fission yeast. In The Molecular and Cellular Biology of the Yeast Saccharomyces: Cell
Cycle and Cell Biology (J. Pringle, J. Broach,
and E. W. Jones, eds.) pp. 697-763. Cold Spring
Harbor Laboratory Press, Cold Spring Harbor,
N.Y.
Moreno, S., Hayles, J., and Nurse, P. 1989. Regulation of p34cdc2 protein kinase during mitosis.
Cell 58:361-372.
Moreno, S., Klar, A., and Nurse, P. 1991. Molecular
genetic analysis of the fission yeast Schizosaccharomyces pombe. Meth. Enzymol. 194:795823.
Moser, B.A. and Russell, P. 2000. Cell cycle regulation in Schizosaccharomyces pombe. Curr.
Opin. in Microbiol. 3:631-636.
Nasmyth, K. 1977. Temperature-sensitive lethal
mutants in the structural gene for DNA ligase in
the yeast Schizosaccharomyces pombe. Cell
12:1109-1120.
Russell, P. and Nurse, P. 1986. cdc25+functions as
an inducer in the mitotic control of fission yeast.
Cell 45:145-153.
Umesono, K., Toda, T., Hayashi, S., and Yanagida,
M. 1983. Two cell division cycle genes NDA2
and NDA3 of the fission yeast Schizosaccharomyces pombe control microtubular organisation and sensitivity to anti-mitotic benzimidazole compounds. J. Molec. Biol. 168:271-284.
Verkade, H.M. and O’Connell, M.J. 1998. Cut5 is a
component of the UV-responsive DNA damage
checkpoint in fission yeast. Mol. Gen. Genet.
260:426-433.
Willer, M., Hoffmann, L., Styrkarsdottir, U., Egel,
R., Davey, J., and Nielsen, O. 1995. Two-step
activation of meiosis by the mat1 locus in Schizosaccharomyces pombe. M o l Cell Bio l
15:4964-4670.
Yamamoto, M. 1996. The molecular control mechanisms of meiosis in fission yeast. Trends Biochem. Sci. 21:18-22.
Contributed by Susan L. Forsburg
The Salk Institute for Biological Studies
La Jolla, California
Yeast
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Introduction of DNA into S. pombe Cells
UNIT 13.17
Transformation is the essential method by which DNA is introduced into a cell, whether
it is a plasmid episome or an integration construct. Fission yeast is readily transformed
using a variety of methods, but efficiencies, expressed as transformed colonies per
microgram plasmid, are generally lower than S. cerevisiae (albeit still more than sufficient
for plasmid recovery, screening of plasmid libraries, repair of gapped plasmids, and other
standard yeast procedures). There are several possible explanations. First, the protocols
have not been as carefully optimized. Second, maintenance of episomes may be affected
by performance of the replication origin, and the absence of a centromere or other
segregation method. Third, mutants affecting DNA dynamics may be harder to transform.
S. pombe is efficient at homologous integration of fragments of DNA, although it is also
adept at nonhomologous recombination. For maximum efficiency of integration and
minimum background, tracts of homology >300 base pairs are recommended; targeted
integration occurs with smaller homology tracts (e.g., 50 base pairs), but generally have
substantial background of nonhomologous events which must be screened out (Bähler et
al., 1998; Grallert et al., 1993; Keeney and Boeke, 1994). Most plasmids are selected
using nutritional markers on EMM plates. However, G418 resistance requires selection
on YES plates.
S. pombe can be transformed by lithium acetate (see Basic Protocol and Alternate Protocol
1), electroporation (see Alternate Protocol 2), or protoplast treatment (see Alternate
Protocol 3). In addition, protoplasts can be induced to fuse and undergo karyogamy, which
provides a means of mating sterile strains (see Support Protocol).
NOTE: All solutions and equipment coming into contact with cells must be sterile, and
proper sterile technique should be used accordingly.
TRANSFORMATION USING LITHIUM ACETATE
The principles of this method (Okazaki et al., 1990) are similar to S. cerevisiae, although
maximum efficiency requires that the lithium acetate have a lower pH. Reported transformation frequency is up to 105 transformants per microgram DNA. This is a laborious
but high-efficiency method; a more rapid version with somewhat reduced efficiency
follows (see Alternate Protocol 1).
BASIC
PROTOCOL
Materials
YES medium (UNIT 13.15)
S. pombe
MB medium (see recipe)
0.1 M lithium acetate (adjust to pH 4.9 with acetic acid)
Plasmid DNA
TE, pH 7.5 (APPENDIX 2)
50% (w/v) PEG 4000, 25° or 30°C
50% YE medium (see recipe for YES in UNIT 13.15)
Selective EMM plates (UNITS 13.15)
25° or 32°C incubators
1-liter flask
43°C water bath
Additional reagents and equipment for culturing yeast (UNIT 13.7)
NOTE: If necessary, a 30° rather than a 32°C incubator can be used.
Yeast
Contributed by Susan L. Forsburg
Current Protocols in Molecular Biology (2003) 13.17.1-13.17.8
Copyright © 2003 by John Wiley & Sons, Inc.
13.17.1
Supplement 64
Prepare cells for transformation
1. Two days before the experiment, inoculate 5 ml YES medium with a single S. pombe
colony of the strain to be transformed. Grow overnight to saturation at 32°C.
The saturated overnight culture may, if desired, be prepared up to several days in advance
of the transformation and stored at 4°C. EMM (UNIT 13.15) should be used if selection is
required.
2. The night before transformation, inoculate a 1-liter sterile flask containing 150 ml
MB medium with an appropriate amount of the saturated culture and grow overnight
at 32°C to an OD600 of 0.5. Calculate the volume of starter culture from which to
generate a larger overnight culture which is grown up with shaking.
Y × OD f ) / 2n
(
vol. inoculated (ml) =
OD
where Y is the volume of overnight culture required, n is the expected number of
generations in this media during the expected grown period, ODf refers to the desired
OD for the exponential culture (generally 0.5), and OD refers to the OD600 of the
starter culture.
The expected number of generations when inoculating from a saturated starter culture must
take into account a 1-generation lag phase to allow the cells to exit stationary phase and
re-enter the cell cycle. For example, for a 12-hr growth period of a strain with a 3-hr
doubling time, the total number of possible growth generations is 12/3 = 4. If diluting from
an already growing exponential culture, then n = 4. However, if inoculating from a
stationary starter culture, there is one lag generation, so the actual growth generations =
4 − 1, or 3.
The amount of culture to be inoculated depends upon the growth rate of the strain which
is influenced by growth temperature, genotype, age of the starter culture, and growth media.
A healthy wild-type strain in complete medium (YES or EMM) at 32°C has a generation
time of between 2.5 and 3 hr during exponential growth. MB is a stringent minimal medium
that has severely reduced growth rate compared to normal EMM; generation time will be
≥5 hr. It is not clear why this medium improves efficiency of transformation, but it may
affect cell wall permeability.
Strains with particular auxotrophic markers or other mutations may be unable to grow in
MB sufficiently well to generate a large working culture. In this case, cells may be grown
to mid-exponential phase in EMM (OD600 = 0.5) and diluted into MB for two additional
generations to gain some of the benefits of growth in MB. Low-glucose EMM can be used
in place of MB, but transformation efficiency may be reduced. How much depends upon
the individual strain.
3. Harvest the cells 5 min at 228 × g (3000 rpm), room temperature. Wash cells in 40
ml sterile water and centrifuge again.
Transform with lithium acetate
4. Resuspend the cells at 1 × 109 cells/ml (0.5 to 1.5 ml) in 0.1 M lithium acetate, pH
4.9, and dispense 100-µl aliquots into microcentrifuge tubes. Incubate 60 min at
32°C, or 25°C for temperature-sensitive (ts) mutants.
Cells will sediment at this stage. Cells can be left for up to 120 min without harm. Each
tube is sufficient for a single transformation.
5. Add 1 µg plasmid DNA in 15 µl TE, pH 7.5 to each tube. Mix by gentle vortexing,
completely resuspending cells that sedimented during the incubation. Do not allow
the tubes to cool down.
Introduction of
DNA into S.
pombe Cells
Duplicate transformations using the same plasmid for each tube maximizes total transformants (e.g., screening a library).
13.17.2
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Current Protocols in Molecular Biology
6. Add 290 µl of 50% (w/v) PEG 4000, 32°C (25°C for ts mutants) to each tube. Mix
by gentle vortexing and incubate 60 min at 32°C (25°C for ts mutants).
7. Heat shock 15 min in a 43°C water bath or heat block. Cool the tubes and incubate
10 min at room temperature.
8. Microcentrifuge 2 min at 5000 rpm. Carefully remove the supernatant by aspiration.
9. Resuspend each tube in 1 ml of 50% YE medium by pipetting up and down with a
1000-µl micropipettor.
10. Transfer each tube suspension to a 50-ml flask and dilute with 9 ml of 50% YE.
Incubate with shaking at least 60 min at 32°C (25°C for ts mutants).
11. Plate <0.3-ml aliquots onto EMM plates selective for the marker of interest at 32°C
(25°C for ts strains). If necessary, centrifuge the cells from a single transformation
and resuspend in 1 ml EMM medium to spread more cells on a plate.
Colonies will generally form in 3 days at 32°C for wild-type cells
RAPID LITHIUM ACETATE PROCEDURE
This rapid method (Kanter-Smoler et al., 1994) reports efficiencies of up to 2 × 10 per
microgram plasmid DNA. It uses small cultures and does not include a grow-out period
for phenotypic expression.
4
ALTERNATE
PROTOCOL 1
Additional Materials (also see Basic Protocol)
Low-glucose EMM with appropriate supplements (UNIT 13.15)
100 mM lithium acetate/1mM EDTA (pH 4.9)
40% (w/v) PEG 3350/100 mM lithium acetate/1mM EDTA, pH 4.9
42°C water bath
Additional reagents and equipment for growth of yeast (UNIT 13.7)
1. Grow a 10-ml culture of S. pombe in low-glucose EMM low to a density of 0.5–1 ×
107 cells/ml (OD600 = 0.2 to 0.5) as described above.
2. Wash once with 10 ml sterile water and resuspend in 1 ml sterile water. Transfer to
a sterile 1.5-ml microcentrifuge tube and pellet cells by microcentrifuging briefly at
20°C.
3. Wash once with 0.2 ml of 100 mM lithium acetate/1mM EDTA, pH 4.9, and resuspend
in 50 µl lithium acetate/EDTA.
4. Add 1 µg DNA in up to 30 µl water or TE, pH 7.5, and add 300 µl of 40% (w/v) PEG
3350/100 mM lithium acetate/1mM EDTA, pH 4.9.
5. Incubate 30 min with agitation at 32°C.
6. Heat shock 15 min in a 43°C water bath without agitation.
7. Pellet by microcentrifuging briefly at 20°C. Resuspend in 1 ml TE, pH 7.5, and plate
200-µl aliquots on EMM plates selective for the plasmid marker.
Colonies will generally form in 3 days at 32°C for wild-type cells.
Yeast
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Supplement 64
ALTERNATE
PROTOCOL 2
ELECTROPORATION OF S. POMBE CELLS
ALTERNATE
PROTOCOL 3
PROTOPLAST PROCEDURE
The electroporation protocol for S. pombe is similar to that for S. cerevisiae (see UNIT 13.7);
however, it is not necessary to plate the S. pombe cells on sorbitol-containing selection
plates and in fact, doing so will actually retard growth. The sorbitol in which cells are
suspended following treatment is sufficient for osmotic protection.
This is a time-consuming protocol that is seldom used any more, although it can be useful
for creating diploids between nonmating strains (see Support Protocol for protoplast
fusion). Transformation frequency is ∼1 × 104 to 5 × 104 transformants per microgram
DNA.
Additional Materials (also see Basic Protocol)
EMM medium (UNIT 13.15)
Cit/phos/EDTA (see recipe)
Cit/phos/sorbitol (see recipe)
NovoZym 234 (BiosPacific)
10 mM Tris⋅Cl, pH 7.6 (APPENDIX 2)/1.2 M sorbitol
10 mM Tris⋅Cl, pH 7.6 (APPENDIX 2)/10 mM CaCl2/1.2 M sorbitol
10 mM Tris⋅Cl, pH 7.6 (APPENDIX 2)/10 mM CaCl2/20% (w/v) PEG 4000
CaCl2/sorb/YE (see recipe)
Selective EMM sorbitol plates (UNIT 13.15)
50-ml plastic centrifuge tube
37°C incubator
29° to 32°C incubator
Additional reagents and equipment for counting cells using a hemacytometer
(APPENDIX 3F)
Prepare cultures
1. Grow a 200-ml culture of S. pombe to an OD600 of 0.2 to 0.5 (up to 1 × 107 cells/ml)
in EMM medium.
2. Harvest cells by centrifuging 5 to 10 min at 4000 × g (5000 rpm), room temperature.
Decant the supernatant and resuspend the pellet in 10 ml cit/phos/EDTA. Transfer to
50-ml plastic centrifuge tube.
3. Harvest cells and resuspend each tube in 5 ml cit/phos/sorbitol.
Form protoplasts
4. Add 25 mg NovoZym 234. Immediately incubate 15 to 30 min at 37°C until
spheroplasts have formed (check that cells have rounded up under a microscope).
Using different enzymes (e.g., Sigma’s lysing enzyme) changes the shape of the cells when
protoplasting. This is because different enzymes break different sugar linkages in the cell
wall. Depending upon which linkages are broken, sometimes the cells get round, sometimes
they stay rodlike, and sometimes they ooze out of their jackets like bright little refractile
balloons, leaving ghosts behind. It is possible to check protoplasting efficiency for any
morphology by taking a small aliquot (a few microliters) and dropping them on a slide with
a large drop of 0.5% to 1% SDS. Protoplasted pombe will lyse in these conditions.
From this point, the protoplasts are extremely fragile, so handle gently.
Introduction of
DNA into S.
pombe Cells
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Current Protocols in Molecular Biology
5. Add 35 ml of 10 mM Tris⋅Cl, pH 7.6/1.2 M sorbitol and divide between 2 to 4 tubes
(no more than 3 × 108 spheroplasts/tube). Centrifuge 5 min at 1000 × g (2000 rpm),
20°C.
It is better to centrifuge at slow speed and repeat than to pellet too vigorously.
6. Resuspend gently in 1 ml Tris⋅Cl/sorbitol, then add an additional 20 ml and invert
several times to wash. Harvest and repeat. After the last resuspension in 1 ml, take a
sample and count the number of protoplasts using a hemocytometer (APPENDIX 3F).
7. Resuspend at 2 to 5 × 108 protoplasts/ml in 10 mM Tris⋅Cl, pH 7.6/10 mM CaCl2/1.2
M sorbitol and pool the contents of the tubes.
Transform protoplasts
8. Add 1 to 10 µg plasmid in no more than 10 µl water to 100 µl protoplasts in a
microcentrifuge tube. Incubate at room temperature for 15 min.
9. Add 1 ml of 10 mM Tris⋅Cl, pH 7.6/10 mM CaCl2/20% (w/v) PEG 4000 and incubate
15 min at room temperature.
10. Microcentrifuge briefly at 20°C. Drain well and resuspend the protoplasts in 0.2 to
0.5 ml CaCl2/sorb/YE. Incubate 30 to 60 min at 30°C.
11. Plate 0.2-ml aliquots onto well-dried, sterile, EMM sorbitol plates. Spread cells very
gently with minimum force (e.g., using a spreader derived from a Pasteur pipet).
Incubate 2 to 5 days at 29° to 32°C (i.e., until transformants appear).
Protoplasts can be aliquotted and stored in 10 mM Tris⋅Cl (pH 7.6)/10 mM CaCl2/1.2 M
sorbitol (see step 7) up to 2 months at −70°C. The frequency of transformation is 1 × 103
transformants per microgram DNA for protoplasts stored in this way.
PROTOPLAST FUSION
This protocol allows creation of a diploid strain using nonmating strains. Cells lacking
cell walls stick together in PEG and may fuse to generate a diploid. Recovery of a diploid
requires selection for complementing markers in the two starting haploids (e.g., ade6M210 and ade6-M216, or ura4+ in one parent and leu1+in the other).
SUPPORT
PROTOCOL
Additional Materials (also see Basic Protocol and Alternate Protocol 3)
Strains with complementary markers
1. Follow protoplast transformation (see Alternate Protocol 3, steps 1 through 6) for
each of two strains with complementing markers.
Complementing mating types are not required.
It is not necessary to count in a hemacytometer.
2. Combine 50 µl of each strain in a sterile tube. Incubate 15 min at room temperature.
3. Add 2 ml of 10 mM Tris⋅Cl (pH 7.6)/10 mM CaCl2/20% (w/v) PEG 4000. Incubate
25 min at room temperature.
4. Plate on EMM sorbitol plates selecting for the complementing markers at the
appropriate temperature (32°C for wild type). As the cells are extremely fragile,
spread gently (e.g., using a spreader from a thin Pasteur pipet).
Growth is reduced on sorbitol plates so it may take a few extra days to see colonies.
Yeast
13.17.5
Current Protocols in Molecular Biology
Supplement 64
REAGENTS AND SOLUTIONS
Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see
APPENDIX 2; for suppliers, see APPENDIX 4.
CaCl2/sorb/YE
10 mM CaCl2
1.2 M sorbitol
0.5 mg/ml yeast extract
5 µg/ml each leu, ura, ade, and his
Sterilize by autoclaving
Store indefinitely at room temperature
Cit/phos/EDTA
2.82 g/liter Na2HPO4
4.2 g/liter citric acid
40 mM EDTA pH 8.0
Sterilize by autoclaving
Store indefinitely at room temperature
Cit/phos/sorbitol
7.1 g/liter Na2HPO4
11.5 g/liter citric acid
1.2 M sorbitol
Adjust to pH 5.6 with 5 M NaOH
Sterilize by autoclaving
Store indefinitely at room temperature
MB medium
0.5 g KH2PO4
0.36 g potassium acetate
0.5 g MgSO4⋅7 H2O
0.1 g NaCl
0.1 g CaCl2⋅2 H2O
5 g (NH4)2SO4
500 µg H3BO4
40 µg CuSO4⋅5 H2O
100 µg KI
200 µg FeCl3⋅6 H2O
400 µg MnSO4⋅H2O
200 µg Na2MoO4⋅2 H2O
400 µg ZnSO4⋅7 H2O
5 g glucose
10 µg biotin
1 mg calcium pantothenate
10 mg nicotinic acid
10 mg myo-inositol
150 mg uracil for ura4− strains or leucine for leu1− strains
Adjust volume to 1 liter with water
Sterilize by autoclaving
Store indefinitely at room temperature
Introduction of
DNA into S.
pombe Cells
13.17.6
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Current Protocols in Molecular Biology
COMMENTARY
Background Information
Critical Parameters
Early attempts to transform fission yeast
relied upon S. cerevisiae plasmids and methods
(Beach and Nurse, 1981). Because most S.
cerevisiae plasmids and markers are poorly
maintained in S. pombe, cloning of S. pombe
specific origins and auxotrophic markers significantly improved efficiencies (for review,
Russell, 1989, and Siam et al., 2003). While
standard S. cerevisiae protocols, most notably
electroporation, work to some extent in S.
pombe, better success with the protoplast and
lithium acetate methods is generally observed
using S. pombe–specific methods.
The choice of method is determined by the
equipment and time available. For example,
electroporation (see Alternate Protocol 2) is
extremely fast and simple to perform, but requires access to an electroporator. Recent studies report improving efficiency following
treatment with DTT (Suga and Hatakeyama,
2001) and also describe storage of frozen electrocompetent cells (Suga and Hatakeyama,
2003).
The classic lithium acetate method (see Basic Protocol) is the most efficient technique, but
it is also quite time consuming and relies on
growth in a stringent minimal medium, as well
as a grow-out period for phenotypic expression.
A faster method should be suitable for most
routine transformations (see Alternate Protocol
1). In addition, lithium acetate has been reported to be more efficient than protoplast treatment (see Alternate Protocol 2) for recovery of
homologous integrants (Grallert et al., 1993).
Methods have also been reported for preservation of frozen competent cells using this
method (Broker, 1993).
Protoplast preparation was the first transformation technique reported for fission yeast
(Beach and Nurse, 1981), but has fallen out of
favor with the development of other methods.
Homologous integration is less effective using
this method than lithium acetate, with events
skewed towards nonhomologous integration
(Grallert et al., 1993), yet protoplast fusion
mediated by PEG allows recovery of diploids
from strains that are otherwise unable to mate
(see Alternate Protocol 3 and Support Protocol). Competent protoplasts can also be preserved in the freezer (Jimenez, 1991).
Further information about transformation
methods may be found in Moreno et al. (1991),
Alfa et al. (1993), and Gould (2003).
The most important variable of any transformation procedure is the growth of the target
strain. For all methods, cells in midexponential
phase are preferred. Growth in low glucose
medium has been reported to enhance transformation efficiency for lithium acetate (KanterSmoler et al., 1994; Okazaki et al., 1990), but
has not been tested for electroporation. Transformation of mutant strains, especially those
defective in genes affecting DNA replication or
repair, or temperature-sensitive strains, may
have reduced efficiency.
The plasmid or other DNA being used
should be of high quality, as nonspecific inhibitors of transformation may be present in crude
preparations. The number of transformants obtained per microgram generally decreases with
larger amounts of DNA, although total transformants will still increase. Especially for electroporation, DNA should be in a small volume
without any excess salt.
Targeted integration of linear plasmids or
fragments varies enormously in efficiency, depending upon the actual locus and the amount
of homology used to target the event (Grallert
et al., 1993; Keeney and Boeke, 1994). A high
background of nonhomologous integrants must
often be screened to find the desired clone.
Increasing the homology generally results in
much more efficient targeting. Protoplast transformation is not recommended for homologous
integration (Grallert et al., 1993).
Troubleshooting
Transformation is highly variable depending upon the strain, plasmid, and conditions of
growth. Mutants defective in DNA replication
are particularly difficult to transform, and may
be slow to form colonies; however, simply
transforming a known plasmid into a strain
does not require high efficiency and can be
performed without much concern or effort. In
contrast, transformation of a plasmid library
relies upon efficient recovery of a large number
of colonies. In this case, a control plasmid
containing the marker of interest should be used
to optimize conditions, including temperature
of growth before and after transformation,
growth stage of the culture (mid- versus late
exponential), and method employed.
A special consideration is transformation
with a library to complement a temperaturesensitive mutant. Failure to recover any transYeast
13.17.7
Current Protocols in Molecular Biology
Supplement 64
formants at the restrictive temperature may
reflect absence of a complementing clone in the
population, or may reflect growth conditions
required to establish the plasmid. If direct plating of the transformation to the restrictive temperature doesn’t work, it often helps to allow
phenotypic expression of the clone at permissive temperature for a day or two before transferring the plates to the restrictive temperature.
In some cases, allowing the transformed colonies to develop at permissive temperature and
then replica plating to restrictive temperature
(e.g., using phloxin B plates to identify the
rescuing clones; see UNIT 13.15) may be the best
way to proceed.
Anticipated Results
Transformation efficiencies may exceed 105
colonies per microgram DNA when the protocols are performed exactly with well-behaved
wild-type strains, but are more likely to be in
the 103 to 104 range with some mutants if frozen
competent cells are used or if short cuts are
taken in the procedures.
Time Considerations
Rapid lithium acetate transformation (see
Alternate Protocol 1) and electroporation (see
Alternate Protocol 2) require minimal manipulation of the target culture: a few washes, treatment with the DNA, and rapid plating. These
can be accomplished within an hour or two. The
classic lithium acetate method (see Basic Protocol) and protoplast preparation (see Alternate
Protocol 3) are significantly more laborious,
including prolonged incubation steps. These
will occupy several hours.
Literature Cited
Alfa, C., Fantes, P., Hyams, J., McLeod, M., and
Warbrick, E. 1993. Experiments With Fission
Yeast. Cold Spring Harbor Laboratory Press,
Cold Spring Harbor, N.Y.
Bähler, J., Wu, J.Q., Longtine, M.S., Shah, N.G.,
McKenzie, A., Steever, A.B., Wach, A.,
Philippsen, P., and Pringle, J.R. 1998. Heterologous modules for efficient and versatile PCRbased gene targeting in Schizosaccharomyces
pombe. Yeast 14:943-951.
Broker, M. 1993. Rapid transformation of cryopreserved competent Schizosaccharomyces pombe
cells. Biotechniques 15:598.
Gould, K. L., ed. 2003. Methods. Academic Press,
San Diego.
Grallert, B., Nurse, P., and Patterson, T.E. 1993. A
study of integrative transformation in Schizosaccharomyces pombe. Mol. Gen. Genet. 238:2632.
Jimenez, J. 1991. Cryopreservation of competent
Schizosaccharomyces pombe protoplasts. Trends
Genet. 7:40.
Kanter-smoler, G., Dahlkvist, A., and Sunnerhagen,
P. 1994. Improved method for rapid transformation of intact Schizosaccharomyces pombe cells.
Biotechniques 16:798.
Keeney, J.B. and Boeke, J.D. 1994. Efficient targeted integration at leu1-32 and ura4-294 in
Schizosaccharomyces pombe.
Genetics
136:849-856.
Moreno, S., Klar, A., and Nurse, P. 1991. Molecular
genetic analysis of the fission yeast Schizosaccharomyces pombe. Meth. Enzymol. 194:795823.
Okazaki, K., Okazaki, N., Kume, K., Jinno, S.,
Tanaka, K., and Okayama, H. 1990. High-frequency transformation method and library
transducing vectors for cloning mammalian
cDNAs by trans-complementation of Schizosaccharomyces pombe. Nucl. Acids Res. 18:64856489.
Russell, P. 1989. Gene cloning and expression in
fission yeast. In Molecular Biology of the Fission Yeast (A. Nasim, P.G. Young, and B.F.
Johnson, eds.) pp. 243-271. Academic Press, San
Diego.
Siam, R., Dolan, W.P., and Forsburg, S.L. 2003.
Choosing and using S. pombe plasmids. Methods In Press.
Suga, M. and Hatakeyama, T. 2001. High efficiency
transformation of Schizosaccharomyces pombe
pretreated with thiol compounds by electroporation. Yeast 18:1015-1021.
Suga, M. and Hatakeyama, T. 2003. High-efficiency
electroporation by freezing intact yeast cells with
addition of calcium. Curr. Genet. 43:206-211.
Contributed by Susan L. Forsburg
The Salk Institute for Biological Studies
La Jolla, California
Beach, D. and Nurse, P. 1981. High frequency transformation of the fission yeast S. pombe. Nature
200:140-142.
Introduction of
DNA into S.
pombe Cells
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