Genetics
YEAST GENETICS
Fred Winston
7.1 Introduction
Key Concepts
• Genetic studies of the yeast Saccharomyces cerevisiae have made monumental
contributions to our knowledge of gene expression and cell growth.
The yeast Saccharomyces cerevisiae is an ideal experimental organism. It is a
microorganism that has a fast rate of growth, with a generation time of only ninety
minutes under optimal conditions. Genetic methods have been developed that allow
straightforward and generally easy manipulation of its genome. Any desired mutation
can be incorporated into the S. cerevisiae genome, allowing powerful genetic
analyses to be performed. S. cerevisiae shares many fundamental properties with
other eukaryotes, including humans. Therefore, what is learned from studies of S.
cerevisiae is often directly relevant to issues in human biology.
Yeast has been the focus of extensive studies in many aspects of molecular biology.
These areas include the cell cycle, recombination, cytoplasmic inheritance, secretion,
transcription, translation, the cytoskeleton, and genomics. Overall, studies in yeast
have made critical contributions to our understanding of cell growth in many ways.
Although S. cerevisiae is the most commonly studied yeast, it is not the only one that
has been used in research. Two other yeasts have also been studied to an extensive
degree. First, research on Schizosaccharomyces pombe has made extremely
important contributions to our understanding of the eukaryotic cell cycle. (See
Genetics 7.25 S. cerevisiae and S. pombe have been invaluable organisms for
elucidating cell cycle control.) Second, the human pathogenic yeast Candida
albicans has also received significant attention from scientists trying to understand
factors that control its virulence in mammalian cells. (See Genetics 7.28 Relevance of
yeast studies to human health.) Most of this chapter discusses studies of S.
cerevisiae, which we will often simply refer to as yeast.
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7.2 A brief history of yeast genetics
Key Terms
The ascus of a fungus contains a tetrad or octad of the (haploid) spores, representing
the products of a single meiosis.
Many of the fundamental genetic facts concerning studies of yeast came from the
work of Øjvind Winge, whose work from the 1930s through the 1950s at the
Carlsberg Laboratory in Denmark provided the critical foundation for modern yeast
studies. Winge developed methods for dissecting tetrads (removing and separating
the four meiotic spores out of the ascus). He also demonstrated Mendelian
segregation in yeast, established that both haploid and diploid states of the organisms
exist, and established that there are two stable mating types.
Another important figure in early yeast genetics, Carl C. Lindegren, published the
first genetic map of yeast in 1949 (2874). Lindegren also helped to lead the yeast
field toward using heterothallic yeast strains (that is, those that are stable for mating
type) for genetic studies.
Genetic research in yeast progressed steadily in the 1960s and 1970s. Mapping
studies demonstrated that the organization of genes in yeast was different from that
of E. coli by showing that operons did not exist. The validity of this idea was proven
as the yeast genome became understood in greater detail; the genome was eventually
completely sequenced in 1996.
Studies in yeast also established that the genetic code was conserved between
prokaryotes and this eukaryote. We now take it for granted that the genetic code is
universal (except in mitochondrial DNA). (See Molecular Biology 7.7 There are
sporadic alterations of the universal code.) However, when yeast was revealed as
using the same nonsense codons as prokaryotes, it was the first time this
conservation was demonstrated outside of the prokaryotic kingdoms (2710).
Studies in yeast exploded in popularity beginning in the late 1970s with the
discovery of methods for the organism's transformation with plasmid DNA (see
Genetics 7.16 Transformation of yeast) and the advent of recombinant DNA
methods. These advances led to the ability to manipulate yeast genes in vivo in ways
never before available in any eukaryote. Over the past twenty years, yeast has
become one of the most intensively and widely studied model systems, analyzed in
over a thousand laboratories worldwide, and making significant contributions in
many different areas.
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Reviews
2874. Lindegren, C. C. (1949). .The yeast cell, its genetics and cytology (St. Louis: Educational
Publishers).
References
2710. Stewart, J. W. and Sherman, F. (1972). Demonstration of UAG as a nonsense codon in bakers'
yeast by amino-acid replacements in iso-1-cytochrome c. J. Mol. Biol. 68, 429-443.
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7.3 Nomenclature for S. cerevisiae genetics
Written studies for any organism rely on a specific nomenclature, which is often
unique for that organism. Such a nomenclature is important because it helps to
communicate information about the nature of mutations and mutant phenotypes. In
this section, we will use standard nomenclature for S. cerevisiae. Examples are
provided in Figure 7.1. A different nomenclature applies for S. pombe, and will be
used in the section on studies of that species.
Figure 7.1 Proteins, mutant
alleles, phenotypes, and genes all
have distinct notations .
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7.4 S. cerevisiae mitotic growth
Key Concepts
• The budding morphology during S. cerevisiae cell division provides an invaluable
tool in cell cycle studies.
Mitosis in S. cerevisiae is similar to that of other eukaryotic cells, with G1, S, G2,
and M phases. S. cerevisiae cells grow mitotically by "budding," as seen in Figure
7.2 and Figure 7.3. At the beginning of the cell cycle (G1), cells have a round or
oval shape. (Diploids are more oval than haploids.) As the cell cycle progresses, a
progeny bud emerges from the parent cell. With the continuation of the cell cycle,
the bud enlarges until finally, late in the cycle, at cytokinesis, the two cells separate.
The progeny cell that forms from the bud is often referred to as the "daughter," and
the parent is called the "mother."
Figure 7.2 Cells at the beginning of the cell cycle (G1) are
unbudded. As cells progress through the cell cycle, a bud
emerges, enlarges, and finally separates, producing a progeny
cell.
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Figure 7.3 S. cerevisiae cells growing mitotically.
This budding process is the same for haploids and diploids. The beginning of G1 in
the mitotic cell cycle is referred to as "START." At this point in the cell cycle, an S.
cerevisiae cell can choose different pathways of growth, including mitosis, meiosis,
or mating, depending on its ploidy, nutritional state, and the proximity of cells of the
opposite mating type.
The mitotic growth by budding has been an extremely useful feature of S. cerevisiae
cells because it allows one to know the stage of the cell cycle by the size of the buds.
Bud size can be combined with a few other easily detectable cell characteristics,
including nuclear division and the morphology of microtubules, to determine
precisely the stage of any S. cerevisiae cell in the cell cycle. S. cerevisiae cell
morphology has been an extremely powerful tool in elucidating fundamental aspects
of the eukaryotic cell cycle. (See Genetics 7.25 S. cerevisiae and S. pombe have been
invaluable organisms for elucidating cell cycle control.)
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7.5 The S. cerevisiae life cycle
Key Concepts
• The ability to grow as a haploid or as a diploid is a key advantage in S. cerevisiae
genetic analysis.
One of the main advantages to genetic studies of S. cerevisiae is that it can grow
mitotically in a stable fashion as either a haploid (with one copy of each
chromosome) or a diploid (with two copies of each chromosome). The great
advantages of this property will be clear in the description of mutant isolation and
analysis. (See Genetics 7.12 Isolation and analysis of yeast mutants—general
approaches.)
Haploids exist in either one of two mating types, called a and α. Mating type is
determined at a genetic locus named MAT that can exist in either of two states, MATa
or MAT α. Haploids of opposite mating type can mate to form diploids. Mating
occurs when cells of the opposite mating type become physically close to each other.
Each haploid cell type secretes a specific mating pheromone that arrests the growth
of cells of the opposite mating type at START of the cell cycle. The cells then fuse,
an event followed by nuclear fusion, resulting in an a/α diploid. (See Molecular
Biology 18 Rearrangement of DNA.)
Diploids are mitotically stable (that is, they will not undergo meiosis) in most growth
conditions. However, when a/α diploids are starved for nitrogen, carbon, or sulfur,
they arrest mitotic growth at G1 and undergo meiosis. Each meiotic diploid gives rise
to four haploid progeny (spores) in a tetrad. This is illustrated in Figure 7.4. (For
additional information, see Molecular Biology 18.2 The mating pathway is triggered
by pheromone-receptor interactions.)
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Figure 7.4 Two haploid cells of the opposite mating type will fuse, producing
a diploid cell. When starved for nitrogen, the diploid will undergo sporulation,
resulting in the formation of a tetrad. A tetrad contains the four products of a
single meiosis.
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Fred Winston
7.6 S. cerevisiae meiotic growth and tetrad analysis
Key Concepts
• Tetrad analysis is a key tool of S. cerevisiae genetics.
Chromosome segregation in S. cerevisiae proceeds similarly to that of other
eukaryotes, with a meiosis I (reductional division) and a meiosis II (equational
division). During meiosis, S. cerevisiae cells undergo dramatic morphogenetic
changes. Each diploid cell that undergoes meiosis produces four haploid progeny as
spores within an ascus, as seen in Figure 7.4. The ascus is also commonly referred to
as a tetrad. The physical association of all of the products of a single meiosis within a
tetrad allows powerful genetic analyses of meiotic progeny in yeast to be performed.
The genetic analysis of the meiotic products within a tetrad requires a method known
as tetrad dissection. Tetrad dissection is the act of separating the four spores of a
tetrad on a solid growth medium. This growth medium allows them to germinate into
mitotically growing cells. For the spores of a tetrad to be separated, the ascus wall
must be partially digested by the enzyme zymolyase. This allows the separation of
the four spores.
After the zymolyase breaks down the ascus wall, tetrad dissection is performed using
a microscope equipped with a device called a micromanipulator. Although several
types of micromanipulators are used in yeast research labs around the world, all of
them employ a very fine glass needle that is used to separate the spores. This appears
in Figure 7.5.
Figure 7.5 The tetrad dissection
microscope enables the researcher
to separate the four spores from the
ascus.
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With the digested tetrads spread onto solid agar medium in a petri plate, the
micromanipulator is used to isolate individual tetrads, separate the spores, and place
each spore into a designated position on the agar. Usually, ten to twenty tetrads are
dissected on a standard-sized petri plate. After dissection, the spores grow into
colonies that can be tested for mutant phenotypes.
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7.7 Using tetrad analysis to determine single-gene
segregation
Key Terms
Sporulation is the generation of a spore by a bacterium (by morphological
conversion) or by a yeast (as the product of meiosis).
Key Concepts
• Single mutations segregate 2:2 within each tetrad.
While tetrad dissection is the act of separating spores from within tetrads, tetrad
analysis involves determining the segregation of genetic markers in the tetrads and
interpreting these segregation patterns. Tetrad analysis allows us to learn several
important types of information about the nature and inheritance of mutations. The
power of tetrad analysis comes from the fact that all four meiotic products are
contained within a single ascus. Mendelian segregation patterns predict specific
patterns of segregation in tetrads, and these can be easily tested by tetrad analysis.
The most fundamental test by tetrad analysis is the one for single-gene segregation.
This determines whether a mutant phenotype is caused by a mutation in a single
gene. If a single gene is the cause, the mutation should segregate in tetrads in the
predicted Mendelian fashion for a single gene.
As an example, consider a mutation in the S. cerevisiaeHIS1 gene, seen in Figure
7.6. This gene is required for histidine biosynthesis; when it is mutant, S. cerevisiae
cells are histidine auxotrophs. An auxotroph is a mutant that is defective for the
synthesis of a metabolite. A prototroph is able to synthesize all required metabolites.
If a his1 mutant is mated by a wild-type HIS1 strain, then the diploids will be
heterozygous for the mutation. After meiosis and sporulation, two spores will
contain the his1 mutation and the other two will contain the wild-type HIS1 gene.
Thus, in each tetrad, two spores will be His+ and two will be His–. This segregation
pattern of 2:2 is diagnostic of the segregation of a single mutation.
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Figure 7.6 A mutation in a single
gene will segregate in a Mendelian
manner. Two progeny will carry the
mutant gene and two will carry the
wild-type gene.
In summary, if a mutant phenotype segregates 2:2, it is caused by a single mutation
in a single gene. However, if it does not segregate 2:2, it is not caused by a single
mutation. As we will see below, other information can be gained from tetrad
analysis.
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7.8 Using tetrad analysis to map genes
Key Terms
A parental ditype (PD) is a tetrad in which two genetic markers are segregating, and
two spores have one parental genotype and two spores have the other parental
genotype.
A nonparental ditype (NPD) is a tetrad in which two genetic markers are
segregating, and two spores have one nonparental genotype and two spores have
the other nonparental genotype.
Chromatids are the copies of a chromosome produced by replication. The name is
usually used to describe each of the copies in the period before they separate at
the subsequent cell division.
The four-strand stage of meiosis is the stage after DNA replication, prior to meiosis
I, when the two pairs of sister chromatids are adjacent.
A tetratype (TT) is a tetrad in which two genetic markers are segregating. Two of
the four spores have parental genotypes and two have recombinant genotypes.
Key Concepts
• Tetrad analysis can determine the possible genetic linkage of two genes with
respect to their centromeres and with respect to each other.
• In contrast to the unordered tetrads made by S. cerevisiae, the bread mold
Neurospora crassa produces ordered tetrads that reflect the positions of the
chromatids during meiosis. This order provides an easy way to measure centromere
linkage.
In addition to using tetrad analysis to determine single-gene segregation, we can also
use it to measure the genetic map position of a gene. This can be either relative to
another gene (two-gene segregation), or relative to its centromere (gene-centromere
segregation).
The ability to determine the genetic map position of a gene is often critical to its
study. Even now, when the genome of S. cerevisiae has already been sequenced, S.
cerevisiae geneticists are frequently in the position of studying mutations whose
identities are unknown. Often, the gene's identity is determined by cloning (see
Genetics 7.17 Isolation of S. cerevisiae genes by cloning).
However, in some cases, cloning is difficult or is not performed at an early stage of
mutant analysis because of the large number of mutations being studied. In these
instances, tetrad analysis is the key. Also, as described in the cloning section, tetrad
analysis is an important part of analyzing a cloned gene. Furthermore, studies of
other yeasts and fungi, whose genomes are not yet sequenced, rely on tetrad analysis
for genetic analysis. We will learn about using tetrads to help determine map position
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in another fungus, Neurospora crassa. (See Genetics 7.11 Ordered tetrads of
Neurospora offer special advantages.)
The ability to use tetrad analysis to map genes derives directly from the special
information that we obtain from having all four meiotic products in an ascus. As
described below, this allows us to make and test specific predictions regarding the
segregation patterns that will occur depending on whether two genes are linked or
unlinked.
In addition, tetrad analysis allows the use of the centromere as a genetic marker. This
is because the centromeres of nonsister chromatids always segregate away from each
other in meiosis I, and the centromeres of sister chromatids always segregate away
from each other in meiosis II. These aspects of tetrad analysis are further described
below.
To understand how the analysis of gene-gene linkage and of gene-centromere linkage
is performed, we must first define the three types of tetrads that can arise when two
genes are segregating during meiosis.
First, let us consider a genetic cross in which we will follow mutations in two genes
that are on different chromosomes, that is, two genes that are completely unlinked to
each other. The two mutations we will follow will be leu2, on chromosome III, and
trp1, on chromosome IV ( Figure 7.7).
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Figure 7.7 The three types of tetrad produced by random segregation of two
markers are: (1) parental ditypes (PD); (2) nonparental ditypes (NPD); and (3)
tetratypes (TT).
A cross between these two mutants will involve parents of the following genotypes:
MATaleu2 and MATα trp1. Therefore, the MATa parent is a leucine auxotroph (Leu–)
and the MATα parent is a tryptophan auxotroph (Trp–). To perform the cross, the two
parents are mixed together by mixing colonies of each strain together with a sterile
toothpick on solid growth medium that contains all required nutrients. In this way,
the two haploid parents can mate, and LEU2/leu2TRP1/trp1 heterozygous diploids
will form. After diploids form, they are purified, either by selecting for the
prototrophic diploids or by screening for the unique morphology of the diploid
zygote shortly after the cells fuse. In the example given, diploids can be selected by
growing the mating mixture on medium that lacks both leucine and tryptophan. The
diploid will be a prototroph (Leu+ Trp+).
The diploids are then sporulated by their transfer to medium that imposes starvation
for nitrogen. After sporulation, the tetrads are dissected and analyzed to determine
the segregation of the leu2 and trp1 mutations.
Analysis of this cross shows that three types of tetrads arise as a result of the normal
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segregation of the markers, as pictured in Figure 7.7. To understand two of these
types of tetrads, let us first consider what happens in the absence of any
recombination. The first class of tetrad is that in which the two parental sets of
markers segregate away from each other at meiosis I. The resulting tetrads have two
spores bearing the genotype of one parent (leu2TRP1) and the other two spores
bearing the genotype of the other parent (LEU2trp1). This class of tetrad, having the
two classes of parental spores, is called a parental ditype (PD).
The second class of tetrad is that in which segregation at meiosis I occurred in the
opposite fashion. The result is a tetrad with two different classes of spores,
LEU2TRP1 and leu2trp1. This class of tetrad, having two classes of nonparental
spores, is called a nonparental ditype (NPD).
Because segregation at meiosis I is random and occurs at equal frequency (in
accordance with Mendel's First Law), for two unlinked genes, the frequency of PD
tetrads equals the frequency of NPD tetrads. This prediction constitutes one of the
fundamental tests of genetic linkage in yeast. Therefore, when testing the possible
linkage of two mutations with unknown map positions, the first question to answer
is, "Does the number of PDs equal the number of NPDs?" If the answer is yes, the
mutations (and hence the genes) are unlinked. If the answer is no, there is linkage.
(For the method of analyzing this linkage, see Genetics 7.10 Using tetrad analysis to
determine gene-gene linkage.)
The third type of tetrad arises as the result of a crossing over, or recombination,
between one of the genes and its centromere. Crossing over during yeast meiosis
occurs after replication, when there are four chromatids. This stage is referred to as
the four-strand stage, with the word strand referring to a chromatid.
Recombination between a marker and its centromere at this stage will result in a
tetrad that contains four spores, each with a distinct genotype. In our example of the
unlinked LEU2 and TRP1 genes, the four types of spores will be:
• LEU2TRP1,
• leu2trp1,
• LEU2trp1, and
• leu2TRP1.
This class of tetrad, having four classes of spores, is called a tetratype (TT).
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7.9 Calculating gene-centromere linkage from tetrad
frequencies
For two unlinked genes (where PD = NPD), a TT tetrad arises as the result of a
crossover between one of the markers and its centromere. (We will see later that TT
tetrads can also arise in another way.) Thus, for two unlinked genes, the frequency of
TT tetrads depends on the linkage of each gene to its centromere. This property can
be used to determine whether a gene is linked to its centromere or not.
Let us consider first the case in which both genes are very tightly linked to their
centromeres. One of the markers that we have already encountered, trp1, is very
tightly centromere-linked and is on chromosome IV. A second centromere-linked
marker is met14, on chromosome XI. Since these two genes are on different
chromosomes, they are not linked to each other. Therefore, in a cross of trp1 by
met14, tetrad analysis will show that PD = NPD. However, because there will be
very little recombination between either gene and its centromere, the frequency of
TT tetrads will be extremely low.
By determining the frequency of TT tetrads, the possible linkage of any marker to its
centromere can be determined by a genetic cross. In such a cross, the marker of
interest is crossed by a known marker that has been determined to be unlinked to the
marker of interest and that also is known to be tightly centromere-linked (such as
trp1). Thus, in a cross of this kind, trp1 serves as a genetic marker for a centromere.
(In this case, it is the centromere on chromosome IV, but a centromere-linked marker
on any chromosome works equally well.) In this type of cross, the frequency of TT
tetrads can be used to calculate the linkage between a marker and its centromere. For
example, the TRP1 gene is approximately one map unit from its centromere,
meaning that only one percent of the tetrads will have a recombination event between
trp1 and its centromere. Therefore, the TT tetrads in this cross will arise virtually
entirely by crossovers between the unknown marker and its centromere, as shown for
leu2 in Figure 7.7.
To understand how tetrad data can be used to measure gene-centromere linkage, we
will consider a cross between trp1 and a hypothetical new mutation called new1.
From a cross of trp1 by new1, we obtain the following numbers of tetrads: 40 PD; 40
NPD; and 20 TT.
What have we learned about the possible centromere linkage of new1 from these
data? First, because PD = NPD, we know that trp1 and new1 are unlinked to each
other. What about new1-centromere linkage?
Recall that linkage is determined by the number of recombinants divided by the
number of total progeny. (See Molecular Biology Supplement 3 Linkage and
mapping.) In our example, there are 20 TT tetrads. Because each tetrad has two
recombinant spores, there are a total of 40 recombinant spores. The total progeny
equals 400 spores (from the 100 total tetrads). Therefore, the linkage is calculated by
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40 recombinants/400 total progeny = 10 map units. As a result, new1 is 10 map units,
or centimorgans, from its centromere.
This mapping function for yeast is usually expressed by an equivalent function
expressed in terms of tetrads: linkage = ½ TT/total tetrads (PD + NPD + TT). As an
example of a real centromere-linked marker, we can also consider the data compiled
for the case of LEU2. In data compiled from many crosses that examined segregation
of LEU2 in relation to a tightly centromere-linked marker, there were 404 TT tetrads
out of a total of 4,196. This equals 4.8 map units between LEU2 and its centromere
on chromosome III.
What do we expect when a gene is unlinked to its centromere? In this case, there is a
predicted ratio of PD:NPD:TT of 1:1:4. To understand how this ratio arises, let us
consider a second cross where the second marker is unlinked to its centromere. In
this case, if we consider all possible patterns of gene segregation to occur at equal
frequency, we see that six classes of tetrads can form at equal frequency with respect
to segregation of two unlinked markers. As Figure 7.8 shows, one class is a PD, a
second class is an NPD, and the remaining four classes are TT tetrads, resulting in
the 1:1:4 ratio.
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Figure 7.8 The random segregation of two markers results in a PD:NPD:TT ratio of
1:1:4. One possible segregation pattern is shown for the first marker, trp1. The
segregation of the second marker, his3, is shown in all possible configurations with
respect to trp1. These ratios of PD:NPD:TT will always be 1:1:4 for all possible
configurations of trp1 segregation.
Note, then, that we can deduce centromere linkage for any two markers that are
unlinked to each other. That is, for any cross involving a known centromere-linked
gene and a gene whose position is unknown, we can ask the general question, "Is the
ratio 1:1:<4?" If the answer is yes, then the gene is centromere-linked. A calculation,
using the function linkage = ½ TT/total tetrads, as just described, is required to
determine the actual linkage. If the ratio is 1:1:4, then the gene is unlinked to its
centromere.
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7.10 Using tetrad analysis to determine gene-gene
linkage
Now let us consider a different type of linkage, in which two genes are linked to each
other. By tetrad analysis we can determine the degree of genetic linkage between
such markers. In the extreme cases of two genes that are completely linked, with no
detectable recombination between them, we expect that tetrad analysis will reveal
that all tetrads are PD tetrads. What about cases where the two genes are linked but
there is some recombination between them?
In those cases, the number of PD tetrads will be greater than the number of NPD
tetrads (PD > NPD). This is because PD tetrads arise from no recombination. In
contrast, for two linked genes, NPD tetrads arise by a double crossover between the
two linked markers. As for centromere linkage, in the case of closely linked genes,
we can calculate the degree of linkage based on the frequencies of the different types
of tetrads produced.
Recombination between linked genes can actually produce PD, NPD, and TT tetrads.
First, if there is a single crossover between the two markers, a TT tetrad is the
product. Recall that for two unlinked genes, when PD = NPD, TT tetrads can arise by
a crossover between a gene and its centromere. However, in the case of linkage,
when PD>NPD, TT tetrads arise by a single crossover between the two genes, as in
Figure 7.9.
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Figure 7.9 A single crossover between
linked markers produces a TT tetrad.
In theory, we can calculate the linkage between two genes as we did for
gene-centromere linkage, using the frequency of TT tetrads. However, recombination
in yeast occurs at a high enough frequency that double crossovers occur and
contribute to the observed linkage. Using tetrad analysis, we can measure the
frequency of double crossovers and take them into account when calculating
gene-gene linkage.
Remember that meiotic recombination occurs at the four-strand stage. There are four
different classes of double crossovers that occur at a specific frequency between two
linked genes, as we see in Figure 7.10. One of these classes, a "two-strand double,"
produces a PD tetrad; another class, a "four-strand double," produces an NPD tetrad;
and two other classes, both called "three-strand doubles," produce TT tetrads. Thus,
we see that different numbers of crossovers can result in PD and TT tetrads, while
only a double crossover generates an NPD. Therefore, as Figure 7.11 illustrates, the
number of NPD tetrads can be used to measure the frequency of double crossovers.
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Figure 7.10 Double crossovers between linked markers can produce PD, NPD, and
TT tetrads. Shown are the possible configurations of double crossovers when the
first crossover occurs between chromatids 2 and 3.
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Figure 7.11 The three types of tetrads that can arise in a genetic cross. For linked
markers, NPD tetrads can only arise by a double crossover, whereas PD and TT tetrads
can arise in more than one way.
Having now accounted for the classes of tetrads that result from zero, one, and two
crossovers, we can calculate the linkage between two genes. Recall that linkage is
determined by recombinants/total progeny. Therefore, we want to be able to calculate
the total number of recombinants among the tetrads.
There will be two recombinant spores in the tetrads with a single crossover (TT
tetrads), and there will be four recombinant spores in the tetrads with a double
crossover. We can calculate the number of tetrads with double crossovers by
remembering that the NPD tetrads will equal one-fourth of the total number of
tetrads that underwent crossovers. Consequently, the total number of tetrads with
double crossovers equals 4NPD. The tetrads with single crossovers are TT tetrads.
However, TT tetrads can also arise by double crossovers. Because half of the double
crossover tetrads are TT tetrads, that class equals 2NPD. Therefore, the tetrads with
single crossovers equals TT-2NPD. Thus, the total crossovers can be represented by
2(4NPD) + (TT-2NPD) = 6NPD + TT.
The total progeny, represented as tetrads, is equal to PD + NPD + TT. Finally, as we
did for calculating centromere linkage, we multiply the number of progeny by 0.5
because we are expressing linkage in terms of single crossovers within tetrads. As a
result, the formula for gene-gene linkage is
map distance (centimorgans) = [½ (6NPD + TT)/PD + NPD + TT]100
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This is the standard formula used to calculate genetic linkage between two markers
in yeast. Note that the genetic mapping formula for gene-gene linkage will be more
accurate than that for gene-centromere linkage. This is because the former accounts
for both single and double crossovers, while the latter only accounts for single
crossovers.
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YEAST GENETICS
Fred Winston
7.11 Ordered tetrads of Neurospora offer special
advantages
Key Terms
First division segregation (FDS) describes the pattern of meiotic segregation that
occurs when no recombination occurs between a marker and its centromere. In
the absence of recombination, the alleles on the two homologues segregate away
from each other at meiosis I.
Second division segregation (SDS) describes the pattern of meiotic segregation that
occurs when a recombination event has occurred between a marker and its
centromere. In this case, the alleles segregate together at meiosis I and do not
segregate away from each other until meiosis II.
Key Concepts
• In contrast to the unordered tetrads made by S. cerevisiae, the bread mold
Neurospora crassa produces ordered tetrads that reflect the positions of the
chromatids during meiosis. This order provides an easy way to measure centromere
linkage.
In the previous sections, we learned how we can use tetrad analysis in yeast to
determine the position of a gene in relation to its centromere and to other genes. This
type of analysis is also possible in the wide range of fungi that produce their meiotic
products in an ascus. The members of this class are called ascomycetes and include
many yeasts, molds, and mushrooms.
One ascomycete of both great historic and contemporary importance is the bread
mold Neurospora crassa. Studies using this organism led to the one gene-one
enzyme model of Beadle and Tatum in 1941 (1228). Neurospora is the most
intensively studied of the filamentous fungi.
Meiosis in Neurospora produces an ascus that contains eight haploid spores in a
linear array. The steps in meiosis for Neurospora are the same as previously
described for yeast, including meiosis I and meiosis II. However, meiosis II is
followed by an extra mitotic cycle, resulting in the eight spores. The linear order of
the spores occurs because the ascus is narrow, forcing the meiotic spindle to orient
along the length of the ascus during meiosis I and meiosis II. This constraint is
helpful to geneticists because the spores are ordered according to the positions of
their centromeres during meiosis, as pictured in Figure 7.12.
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Figure 7.12 The spores produced by Neurospora, as for yeast, are haploid.
Spores of opposite mating type (A and a) can mate to form a diploid that will
go through meiosis. Following meiosis, a round of mitosis results in 8 spores
in a linear tetrad.
The linear nature of Neurospora tetrads means that it is considerably easier to
determine if a gene is linked to its centromere than it is to do so for yeast. The
linearity allows one to detect recombination between a single marker and its
centromere directly. Recall that in yeast, such an event requires measuring
segregation in relation to a second marker known to be linked to a centromere.
To see how we can detect a crossover between a centromere and a genetic marker in
Neurospora, let us consider Figure 7.13, which shows two alleles of a marker, A and
a. As we follow the segregation of this marker during meiosis, we see that different
tetrad patterns are possible, depending on whether or not recombination occurs
between A/a and its centromere.
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Figure 7.13 Examples of markers that are
unlinked and linked to their centromeres. The
frequencies of the different types of tetrads are
given below each class.
When there is no recombination, the two alleles segregate at meiosis I, resulting in
the 4:4 pattern AAAAaaaa. In this case, the two alleles segregate away from each
other during the first meiotic division, so this segregation pattern is called first
division segregation (FDS).
However, when a crossover occurs between A/a and its centromere, other
segregation patterns form. In the case of a crossover, the A and a do not segregate
away from each other until the second meiotic division. Consequently, these
segregation patterns are called second division segregation (SDS).
It possible to determine the centromere linkage of a gene by measuring the relative
level of FDS and SDS. In the example shown in Figure 7.12 for A/a, the gene is
unlinked to its centromere. Therefore, the frequency of FDS equals the frequency of
SDS.
However, when a gene is centromere-linked, the frequency of SDS will be less. In
the case of an SDS tetrad, half of the spores are recombinant. Thus we can calculate
the centromere linkage with the following formula:
map distance = (½)SDS tetrads/total tetrads × 100
This formula is similar to the one used for calculating centromere linkage in yeast,
with SDS tetrads in Neurospora being equivalent to tetratype tetrads in yeast. An
example of a centromere-linked locus (B/b) is shown in Figure 7.13. In this case, we
get the following:
map distance = (½)(7+8+6+7)/200 × 100 = 14 map units or centimorgans
The measurement of gene-gene linkage in Neurospora is the same as in yeast. We
can determine the frequency of the three possible classes of tetrads—PD, NPD, and
TT—that arise when following the segregation of two genes. The formula for
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calculating gene-gene linkage is the same as described in a previous section for
yeast:
map distance (centimorgans) = [½ (6NPD + TT)/PD + NPD + TT]100
It is worth noting that this formula takes into account the double crossovers, while
the calculation for centromere linkage does not.
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References
1228. Beadle, G. W., and Tatum, E. L. (1941). Genetic control of biochemical reactions in
Neurospora. Proc. Natl. Acad. Sci. USA 27, 499-506.
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YEAST GENETICS
Fred Winston
7.12 Isolation and analysis of yeast
mutants—general approaches
Key Terms
A screen is a search for mutants in which each candidate is tested for the mutant
phenotype of interest.
Selection describes the use of particular conditions to allow the preferential growth
or survival of organisms with a particular phenotype. Selection in the laboratory
is usually strong, while natural selection is usually weak in the short term.
The ability to isolate and analyze mutants affecting important biological processes is
one of the great strengths of yeast genetics. This type of approach has led to the
discovery of many important genes, some of which are conserved in humans. This
kind of research has provided important advances in understanding the control of
gene expression, DNA replication, cell division, secretion, and many other
fundamental processes. In this section, we will go over the basic steps involved in the
classical isolation and analysis of mutations.
To understand any biological process, we can consider the approach of isolating and
analyzing mutants in which that process has gone awry. Then, by analyzing the
mutant phenotypes and the characterization of the genes and gene products
identified, we can learn much about how the process functions in the wild-type state.
This is one of the main goals of genetic analysis in yeast.
When initiating a plan to isolate mutants of interest, it is important to determine if
one should do a screen or a selection for the mutants, as there are potential strengths
and weaknesses to each approach. In a screen, one examines every possible
candidate by one or more phenotypic tests. An example of this would be the
screening of yeast colonies for an amino acid auxotrophy. Using this approach, a
large number of candidates must be directly tested, often requiring extensive effort.
In a large-scale screen of yeast colonies, one can reasonably screen up to
approximately 50,000 colonies. Therefore, the frequency of the desired mutants must
be high enough that, after mutagenesis, they can be found within this total. The
advantage of a screen is that every colony is examined; therefore, mutants with a
range of phenotypes can be discovered.
In a selection, such as one for resistance to a toxic compound, one can identify much
rarer events. Because as many as 107 cells can be spread on a single petri plate that
imposes selective conditions, mutations that arise as infrequently as 1/109 can be
identified by a strong selection. Thus the main advantages of a selection are that it
can detect rare mutants and it is easier to test a very large number of candidates. The
possible disadvantage, however, is that the selection conditions might fail to identify
certain classes of mutants, such as those that might grow poorly. On account of this
possible bias, mutant selection conditions are a matter for careful consideration.
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YEAST GENETICS
Fred Winston
7.13 An example of a genetic screen in yeast
Key Terms
A dominant allele determines the phenotype displayed in a heterozygote with
another (recessive) allele.
A recessive allele is obscured in the phenotype of a heterozygote by the dominant
allele, often due to inactivity or absence of the product of the recessive allele.
A complementation test determines whether two mutations are alleles of the same
gene. It is accomplished by crossing two different recessive mutations that have
the same phenotype and determining whether the wild-type phenotype can be
produced. If so, the mutations are said to complement each other and are probably
not mutations in the same gene.
Interallelic complementation (intragenic complementation) describes the change
in the properties of a heteromultimeric protein brought about by the interaction of
subunits coded by two different mutant alleles; the mixed protein may be more or
less active than the protein consisting of subunits only of one or the other type.
Unlinked noncomplementation (nonallelic noncomplementation) is a situation
where mutations in two different genes fail to complement.
Linkage analysis is a test of position between two different mutants. In yeast,
linkage analysis is done on tetrads, the meiotic progeny of a diploid.
As an example of a mutant hunt, let us consider a screen for yeast mutants that are
unable to use the sugar galactose as a carbon source. This was one of the earliest
classes of mutants extensively studied in yeast by genetic analysis. Early studies
demonstrated that when yeast cells were grown on galactose as a carbon source,
three enzymes required for galactose catabolism were induced. In yeast cells grown
on other carbon sources, these enzymes were not present.
We now know that this regulation occurs through the controlled transcription of the
genes required for galactose catabolism. Among mutants defective for growth on
galactose are classes in which either the genes encoding the catabolic enzymes or
those encoding the regulators of these genes are affected.
We will begin the mutant screen with a haploid strain, as this will allow the direct
identification of recessive mutations. In other eukaryotes that live only as diploids,
such as D. melanogaster and C. elegans, genetic screens for recessive mutations
require crosses to examine progeny in the F1 or F2 generation. These extra crosses
are necessary in order to study progeny in which the mutation has become
homozygous. The ability to isolate mutants directly in yeast haploids greatly
facilitates identifying and analyzing those mutants.
In our screen, shown in Figure 7.14, we will first mutagenize the yeast cells to
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increase the frequency of mutations in the population. Without mutagenesis, the
frequency of spontaneous Gal– mutants would be so low (approximately 10–5 to 10–6)
that they would be difficult to find. The two mutagens most commonly used for yeast
are ethylmethane sulfonate and ultraviolet light, both of which cause primarily single
base pair changes.
Figure 7.14 To screen for mutants, a wild-type yeast
strain is grown up in liquid, diluted to spread
approximately 300 cells/plate, and then mutagenized to
approximately 50 percent survival. The colonies are then
screened by replica plating to test for the mutant
phenotype of interest. In this example, it is an inability to
use galactose as a carbon source.
To begin the mutant screen, then, the wild-type strain will be mutagenized and cells
will be spread on a solid medium that is permissive for the growth of any potential
mutants. In this case, the medium will be one that contains yeast cells' favorite
carbon source, glucose. Within two days, each cell will form a colony that can be
screened for mutant phenotypes.
To identify mutants unable to use galactose as a carbon source, the colonies are
screened by replica plating. In this method, a filter or piece of velvet is pressed
against the original plate, picking up some of each colony. It is then pressed onto a
plate that contains a different medium or on which the organisms will be grown at a
different temperature. Replica plating allows us to compare growth of the same cells
under different conditions, so we can find mutants that survive in one situation but
die in another. In our screen, we replica plate to a solid medium that contains
galactose as the carbon source. Colonies that can grow on glucose but not on
galactose are mutant candidates for defects in galactose catabolism.
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Let us suppose that our mutant screen has worked well and we have identified
several mutants that are unable to grow on galactose as a carbon source. We will
hereafter refer to these as Gal– mutants.
How does a yeast geneticist go about studying many mutants? Four types of genetic
analyses are usually applied to determine the number of genes identified. These tests
are for:
• Single-gene segregation,
• Dominance/recessiveness,
• Complementation, and
• Linkage.
Each of these tests requires forming particular types of diploids using the haploid
Gal– mutants.
First, we can determine whether the Gal– phenotype is caused by a mutation in a
single gene by performing genetic crosses to a wild-type (Gal+) strain. This is
followed by tetrad analysis. Single-gene events will show 2:2 segregation.
Testing whether the mutations are dominant or recessive is easy to accomplish by
mating each Gal– mutant with a wild-type (Gal+) strain, as in Figure 7.15. If the
mutation is recessive, the diploid will be Gal+. If the mutation is dominant, the
mutation will be Gal–. In general, recessive mutations are considered to be loss of
gene function, while dominant mutations are considered to be increase or alteration
of gene function.
Figure 7.15 Tests to see if a mutant is dominant or recessive are done in
diploids. The mutant is mated by a wild-type strain. If the diploid has a
wild-type phenotype, the mutation is said to be recessive. If the diploid has
a mutant phenotype, the mutation is said to be dominant.
To begin to identify how many genes have been identified, we want to perform
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complementation tests. A complementation test is a test of function. Because most
genes express a single function, mutations in different genes are usually in different
complementation groups. Likewise, mutations in the same gene are usually in the
same complementation group. Although exceptions to this pattern occur,
complementation tests are an invaluable aid to determine the number of genes
identified in a mutant hunt and generally to indicate if mutations are in the same or in
different genes.
Complementation tests in yeast are performed in diploid strains made by crossing
one mutant with another one. To construct such diploids, each mutation must be
available in both mating types, a and α. This will be achieved from the crosses
described above that test for 2:2 segregation for the mutations. An example of a
complementation test between different pairs of Gal– mutations is diagrammed in
Figure 7.16.
Figure 7.16 Complementation tests can only be done on recessive mutations. One
mutant is mated by another and the diploid is analyzed. If the diploid has a mutant
phenotype, the mutations fail to complement and are likely in the same gene. If the
diploid has a wild-type phenotype, the mutations complement and are likely in two
different genes.
Let us consider the example of two hypothetical recessive Gal– mutants called gal1
and gal2. To perform a complementation test, we cross an a gal1 mutant by an
α gal2 and isolate a gal1 /gal2 diploid. Then we determine the Gal phenotype of the
diploid. If gal1 and gal2 are mutations in different genes, then the diploid will be
Gal+ because all functions required to grow on galactose will be expressed. However,
if gal1 and gal2 are in the same gene, a function required to grow on galactose will
be lacking and the diploid will be Gal–.
Complementation tests like the one just described are easy to do with yeast on a large
scale. Therefore, it is easy to perform complementation tests on forty or more
mutants to place them into complementation groups. Thus, in general,
complementation tests on recessive mutations constitute the best initial step in
determining the number of genes identified in a hunt for mutants.
However, there are cases when complementation tests will not be useful. For
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example, complementation tests cannot be performed with dominant mutations, as
dominant mutations will confer a mutant phenotype in a heterozygous diploid,
regardless of whether or not the other mutation is in the same gene. In addition,even
with recessive mutations, there are rare cases where mutations in the same gene
complement (called intragenic complementation) or mutations in different genes
fail to complement (called unlinked noncomplementation).
The final type of genetic analysis to perform on the mutants is linkage analysis. A
linkage test is a test of position (in contrast to a complementation test, which is a test
of function). Linkage analysis entails crossing one mutant by another and analyzing
the segregation in tetrads. There are clear expectations for most cases. If two
mutations are in the same gene, they will show very tight linkage, with mostly PD
tetrads as evidence. However, if the two mutations are in different genes, they will
likely be unlinked, resulting in a PD:NPD:TT ratio of 1:1:4.
Because linkage analysis is more time-consuming than complementation analysis, it
is usually performed on a subset of mutants, including representatives of each
complementation group. Linkage tests can help to uncover the rare cases of
intragenic complementation and unlinked noncomplementation.
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YEAST GENETICS
Fred Winston
7.14 Epistasis analysis can identify an ordered
relationship among gene functions
Key Terms
Epistasis analysis usually refers to double mutant analysis to determine possible
order of gene actions. The two mutations must have different phenotypes.
Key Concepts
• Sometimes several genes play distinct roles in a common process.
• Comparison of the single mutant phenotypes with the double mutant phenotype can
sometimes determine their relative order of function.
Sometimes genetic analysis can help us to understand the functional relationships
among different genes that work in a common process. In the case of understanding
the use of galactose as a carbon source, imagine two classes of yeast mutants. The
first class is the one unable to use galactose as a carbon source (Gal–), as described in
our screen above. The second class of mutant, isolated in a separate screen, is quite
distinct. It causes constitutive expression of the enzymes required for galactose
catabolism, even when cells are grown on a different carbon source, such as glucose
(Galc).
What might be the relationship between genes that cause such different phenotypes?
One possibility is that they work in an ordered fashion, perhaps in a pathway. This
possibility can be addressed by a genetic test called epistasis analysis.
Epistasis analysis involves constructing a double mutant and examining its
phenotype. Imagine that the Gal– mutant has lost a regulatory factor that acts
positively (that is, it is required for the expression of the GAL genes), and that the
Galc mutant has lost a repressor of GAL gene expression.
We could do an epistasis test to learn about the genetic relationship of these two
factors by crossing the Gal– mutant by the Galc mutant to create a double mutant.
Then we would determine if the double mutant had a Gal– or a Galc phenotype.
If the double mutant were Gal–, we would say the mutation causing the Gal–
phenotype was epistatic to the mutation causing the Galc phenotype. This would
mean that the mutant gene causing the Gal– phenotype likely functioned more
directly to control GAL gene expression. If the double mutant were Galc, then the
Galc mutation would be epistatic and we would reach the opposite conclusion about
the order of gene function. If the mutants act in different pathways, the phenotype of
the resulting double mutant may be intermediate between the two single mutant
phenotypes, as each exerts its effect independently. In general, epistasis analysis has
been a powerful genetic tool to build models of understanding gene action. We will
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later discuss a related topic, the isolation of extragenic suppressors (see
Genetics 7.21 Suppressor analysis is a proven method to identify interacting genes).
That technique is used to look directly for second mutations that are epistatic to a
beginning mutation of interest.
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YEAST GENETICS
Fred Winston
7.15 The S. cerevisiae genome
Key Concepts
• The S. cerevisiae genome was the first eukaryotic genome to be completely
sequenced.
The S. cerevisiae genome contains 16 chromosomes, ranging in size from
approximately 250 kilobases to 1,500 kilobases. The S. cerevisiae genome was the
first eukaryote genome to be fully sequenced, having been completed in 1996. The
genome is approximately 12,000 kilobases, encoding about 6,000 genes (2708).
The DNA sequence has allowed the unambiguous identification of most S. cerevisiae
genes. The genome sequence has been an invaluable aide in many aspects of S.
cerevisiae studies, particularly in genetic and gene expression studies. Moreover, it
facilitates the cloning of genes (see Genetics 7.17 Isolation of S. cerevisiae genes by
cloning).
S. cerevisiae chromosomes have three classes of elements that are important for their
replication and segregation:
• Each chromosome has several origins of DNA replication. S. cerevisiae origins
of replication were initially identified because of the properties that they
conferred on plasmids (2711). More recently, experiments have demonstrated
that these sequences serve as bona fide origins of DNA replication in the
genome.
• Each S. cerevisiae chromosome has a single centromere. The centromere is the
site of attachment of microtubules and is essential for normal chromosome
segregation. Centromeres in S. cerevisiae were the first functional centromeres
cloned (189).
• At the end of each S. cerevisiae chromosome is a telomere. Telomeres in S.
cerevisiae are similar in structure to those of other fungi and eukaryotes,
consisting of repeated DNA. Genetic experiments in S. cerevisiae have
demonstrated that centromeres and telomeres are required for proper
chromosome stability and segregation.
Using these characterized elements, investigators have constructed recombinant
DNA molecules that can function as chromosomes in S. cerevisiae (592). The
"artificial" chromosomes function in a similar manner to endogenous S. cerevisiae
chromosomes, demonstrating that these three classes of chromosomal elements are
sufficient for chromosome replication and segregation. Using these artificial
chromosomes, investigators have devised screens for mutants that fail to segregate
chromosomes properly during mitosis (2709)(2715). In this way, several factors that
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bind to centromeres and that are required for normal segregation have been
discovered.
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Reviews
189. Clarke, L. and Carbon, J. (1985). The structure and function of yeast centromeres. Annu. Rev.
Genet. 19, 29-56.
2708. Goffeau, A., et al. (1996). Life with 6000 genes. Science 274, 546-563.
References
592. Murray, A., and Szostak, J. W. (1983). Construction of artificial chromosomes in
yeast. Nature 305, 189-193.
2709. Koshland, D., Kent, J. C., and Hartwell, L. H. (1985). Genetic analysis of the mitotic transmission
of minichromosomes. Cell 40, 393-403.
2711. Stinchcomb, D. T., Struhl, K., and Davis, R. W. (1979). Isolation and characterisation of a yeast
chromosomal replicator. Nature 282, 39-43.
2715. Maine, G. T., Sinha, P., Tye, B. K. (1984). Mutants of S. cerevisiae defective in the maintenance
of minichromosomes. Genetics 106, 365-385.
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YEAST GENETICS
Fred Winston
7.16 Transformation of yeast
Key Concepts
• Transformation of yeast allows many types of genetic manipulations that are
impossible using classical genetic approaches.
One of the major technical breakthroughs in the molecular genetic analysis of S.
cerevisiae was the development of the method of transformation of S. cerevisiae with
plasmid DNA (2915). This method opened the door to the cloning of genes and
several other types of genetic manipulations described in the following sections.
Transformation allows the manipulation of genes in vitro, followed by their analysis
in vivo.
Three classes of plasmids are generally useful for transformation of S. cerevisiae.
These three classes are known as:
• Integrating,
• Centromeric, and
• High-copy-number or 2-micron.
Their properties, diagrammed in Figure 7.17, vary in transformation frequency,
stability, and copy number. All three classes of plasmids contain some common
elements:
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Figure 7.17 Three classes of plasmids are most commonly used to transform yeast:
integrating, centromeric, and 2-micron.
• An origin of replication for propagation in E. coli;
• A selectable marker for transformation of E. coli, usually ampicillin resistance;
and
• A selectable marker for transformation of S. cerevisiae, usually a gene such as
the URA3 that complements an auxotrophy. The URA3 encodes a protein
required for pyrimidine biosynthesis.
Use of URA3 as the selectable marker requires the S. cerevisiae strain that will be
transformed possess a ura3 mutation. Transformants, then, are selected on plates
lacking uracil in the media. The common use of URA3 as a marker for
transformation is historical; the URA3 gene was one of the first S. cerevisiae genes
cloned and was therefore available for use in recombinant DNA experiments. Other
genes commonly used as selectable markers in S. cerevisiae plasmids are also genes
that were among the first cloned, including TRP1, HIS3, and LEU2. More recently,
yeast researchers have started to use markers that confer resistance to antibiotics such
as G418. Because wild-type yeast are sensitive to G418, the use of this marker does
not require a particular genotype of the recipient strain.
In addition to transformation using these three types of plasmids, transformation
using linear DNA is also a valuable tool in S. cerevisiae. (For the use of this method
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to construct mutations in genes, see Genetics 7.18 Molecular genetics of S.
cerevisiae ).
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References
2915. Hinnen, A., Hicks, J. B., and Fink, G.R. (1978). Transformation of yeast. Proc. Natl. Acad. Sci.
USA 75, 1929-1933.
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YEAST GENETICS
Fred Winston
7.17 Isolation of S. cerevisiae genes by cloning
Key Terms
To clone a gene is to molecularly identify its DNA sequence. The phrase "to clone"
may also be used to indicate the process of inserting a piece of DNA into a
cloning vector so that multiple copies may be made.
An open reading frame (ORF) is a sequence of DNA consisting of triplets that can
be translated into amino acids starting with an initiation codon and ending with a
termination codon.
Key Concepts
• Cloning of an S. cerevisiae gene is generally performed by complementation of a
mutant phenotype.
• The proof that the correct gene has been cloned requires tetrad analysis.
Once mutations have been analyzed by a combination of complementation and
linkage analysis, the next step is to identify the specific genes that are mutant. This
important goal is achieved by cloning the genes that correspond to the mutations.
The cloning of a gene of interest allows important types of genetic analysis.
The two important reagents for cloning are:
• An S. cerevisiae strain containing the mutation of interest and
• A recombinant DNA library that contains random S. cerevisiae genomic
restriction fragments from a wild-type strain in an appropriate vector.
The S. cerevisiae strain used to clone a gene must contain two particular genetic
markers. First, it must contain the mutation of interest. Because the correct clone will
be detected by complementation, the mutation must be recessive. (The cloning of a
gene corresponding to a dominant mutation is mentioned later in this section.) The
second important marker is one used to select transformants, such as a ura3
mutation.
The recombinant DNA library consists of a collection of plasmids, with each plasmid
containing a random fragment of wild-type S. cerevisiae DNA cloned into a standard
centromeric plasmid vector that can replicate in both E. coli and in S. cerevisiae.
(Cloning a gene for a dominant mutation would require constructing a recombinant
library from the dominant mutant strain.) As mentioned, this marker is often the
URA3 gene. However, other markers are also frequently used.
The S. cerevisiae genomic fragments are generated by partial digestion with a
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restriction enzyme that cuts frequently in genomic DNA. For S. cerevisiae, unlike
larger eukaryotes, introns are rare, and when they are present, they are small.
Therefore, genomic DNA is suitable to make recombinant libraries instead of cDNA.
Each genomic fragment is likely to carry several open reading frames.
As we can see in Figure 7.18, identifying a clone that contains the gene of interest
starts with using the library DNA to transform the S. cerevisiae mutant.
Transformants are identified by the plasmid's selectable marker. In our example,
selection is for Ura+. Then, transformants are screened for those that complement the
phenotype conferred by the mutation of interest. In our example of gal1 mutants, the
Ura+ transformants will be screened for those that are Gal+. This is done by replica
plating to plates that have galactose as the carbon source. These strains are strong
candidates to contain a plasmid clone that contains the wild-type GAL1 gene.
Figure 7.18 Candidate clones are identified after
transformation into the appropriate yeast mutant.
How many transformants need to be screened to identify the desired clone? We can
determine this number by knowing the size of the S. cerevisiae genome
(approximately 12,000 kb) and the average size of the DNA fragment in S. cerevisiae
recombinant plasmid (approximately 15 kb). We use the formula N = ln(1-P)/ln(1-f),
where N equals the number to be screened, P equals the probability of screening the
clone, and f equals the fragment size/genome size. Using this formula, we calculate
that approximately 3,700 transformants are required in order to have a 99 percent
chance of screening every clone. We can easily screen this number of transformants
by replica plating 40 plates with 100 transformant colonies per plate.
Identifying a clone that contains the gene of interest does not yet single out the actual
gene. This is because the average piece of genomic S. cerevisiae DNA in each
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plasmid will be large enough to contain at least two to three genes. The first step to
pinpoint the exact gene of interest on a clone is to determine the precise genomic
segment in the recombinant plasmid. We can ascertain this by sequencing the DNA
at the ends of the insert. Then, by searching a database against the entire yeast
genome sequence, we can identify the exact genomic fragment in the plasmid. In this
way, we reveal the genes present in the insert.
To establish which gene on the plasmid is the one that complements the mutation, we
can use two different methods. First, we can make new plasmids, each containing a
single gene from those on the original plasmid (see Figure 7.19). Only one of these
genes should complement the mutation and identify the gene. The second approach is
to perform classical complementation tests between the mutant of interest and
mutations in each of the candidate genes, if such mutations are available.
Figure 7.19 Subcloning of the open reading frames on the plasmid,
followed by their analysis in the mutant, will identify which open reading
frame complements the mutant phenotype.
One last but important step is needed to verify that the correct gene has been cloned.
The cloning steps described so far have identified the clone candidate based on
complementation, which is a test of function. Just as complementation in diploids
can occasionally be misleading with respect to gene identity, it can be misleading in
cloning as well. This is because at times, a gene on a plasmid will suppress a defect
in a different gene. One possible reason for this situation is that both genes encode
related functions and one function can substitute for the other, especially if it is
expressed at an increased level on the plasmid copy. Therefore, a linkage test—a test
of position—is also necessary to verify that the correct gene has been cloned.
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This test can take many forms, and we will describe one simple version. When the
candidate gene has been identified, a linkage test can be performed between the
original mutation and a known mutation in the gene (identified by cloning), as
illustrated in Figure 7.20. If the two mutations are in the same gene, virtually every
tetrad will be a PD. In contrast, if the gene newly identified by cloning is not the
same as the gene that contains the original mutation, then a different segregation
pattern will be apparent because the two mutations will most likely not be linked. In
this case, PD, NPD, and TT tetrads will appear. The ability to clone a gene and prove
it by these tests leads to the ability to perform more detailed genetic analysis using
molecular genetic techniques, as described in the next section.
Figure 7.20 Verification that the correct clone has been identified is accomplished
by a cross to establish linkage between the original mutation and the gene identified
from the cloning.
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YEAST GENETICS
Fred Winston
7.18 Molecular genetics of S. cerevisiae
Key Terms
A null mutant has a mutation that completely eliminates the function of a gene,
usually because it has been physically deleted.
The polymerase chain reaction (PCR) is a technique in which cycles of
denaturation, annealing with primer, and extension with DNA polymerase are
used to amplify the number of copies of a target DNA sequence by large factors
such as > 106 fold.
An open reading frame (ORF) is a sequence of DNA consisting of triplets that can
be translated into amino acids starting with an initiation codon and ending with a
termination codon.
A conditional-lethal mutation kills a cell or virus under certain (nonpermissive)
conditions, but allows it to survive under other (permissive) conditions.
A temperature-sensitive mutation creates a gene product (usually a protein) that
functions at some low temperature but poorly or not at all at some high
temperature. The converse is a cold-sensitive mutation.
A cold-sensitive mutant is defective at low temperature but functional at normal
temperature.
The plasmid shuffle is a technique used in yeast genetics to screen for mutations in
an essential gene. The technique is based on the fact that, in a strain deleted for an
essential gene, a plasmid bearing a mutant copy of that gene will be able to
replace a plasmid bearing the wild-type gene if the mutant gene is viable under
some conditions.
Gene replacement describes a method in which the version of the gene in the
genome, usually the wild-type gene, is replaced by a mutant form. The
replacement occurs via homologous recombination following transformation.
Key Concepts
• Any mutation made in vitro can be transformed into the S. cerevisiae genome,
replacing the wild-type allele.
With a cloned S. cerevisiae gene in hand, we can perform gene-specific mutagenesis
to analyze gene function in great detail. Three different types of mutageneses can
help to elucidate the role of a gene in vivo. These three mutageneses are:
• Deletion of the gene,
• Random mutagenesis of the gene, and
• Site-specific mutagenesis of the gene.
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A gene is often chosen as the subject of a study based on a particularly interesting
mutant phenotype. For example, one phenotype that has been extremely valuable to
study in S. cerevisiae is that of mutants that arrest growth at particular points in the
cell cycle. These are referred to as cell division cycle or cdc mutants. Such a
phenotype could be caused by very different types of mutations. These could include
one that caused loss of function of the gene product, or one that brought about an
alteration in its normal function.
Therefore, an early step after cloning the gene is to construct a deletion of it,
resulting in a null mutant. Comparison of the phenotype of a null mutant with that
of the original mutant will determine if the original mutant phenotype results from a
loss of function. The analysis will also answer the related issue of whether or not the
gene is essential for viability.
As Figure 7.21 illustrates, such a deletion can be made in a straightforward manner
by using the combination of the polymerase chain reaction (PCR) and S. cerevisiae
transformation.
Figure 7.21 A precise deletion of any gene of yeast
can be made using a DNA fragment synthesized by
PCR.
• The gene encoding a selectable marker is synthesized by PCR, using primers that
contain a short region of homology to the DNA sequences flanking the open
reading frame to be deleted.
• Next, the PCR fragment is used to transform a wild-type diploid yeast strain. A
diploid strain is used in case the deletion to be constructed would cause lethality
in a haploid. In the example shown, transformants are selected by resistance to
the antibiotic G418. The high frequency of homologous recombination in yeast
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allows the limited homology to direct recombination at the correct site.
• The diploid is then sporulated and tetrads are dissected. The marker, G418
resistance, segregates 2:2 and indicates segregation of the deletion mutation. In
the example shown, the deletion of GAL1 allows viability. In the case of the
deletion of an essential gene, only two spores will germinate to grow into
colonies. The two viable spores will never contain the G418 marker. That is,
G418 resistance will be tightly linked to inviability.
Thanks to the knowledge of the complete genome sequence of S. cerevisiae, a
deletion mutation of every gene has now been made (2866). Thus, once a gene is
identified by cloning, an investigator will immediately be able to know its null
phenotype. Because the null phenotype is usually the most extreme phenotype for a
gene, if the null mutant is viable, then it is the mutant form best suited for further
genetic studies.
In addition to a null mutation, there are cases in which the isolation and analysis of
other types of alleles are desirable. One obvious case is when a gene is essential for
viability, because in such a case, the null mutant is inviable. This is generally true for
cdc mutants. For essential genes, then, less severe mutations are required for genetic
analysis.
Random mutagenesis is the best approach for an essential gene that has been
identified as important and worthy of study, but about whose function little else is
known. In such a case, investigators will want to isolate several different mutant
alleles, since different mutations might alter the gene product in different ways.
Observing distinct mutant phenotypes for different alleles of a gene will provide
more information about the role of the gene product. A general method to isolate
multiple alleles of a gene is described in this section.
One particularly valuable class of mutation in an essential gene is a
conditional-lethal mutation. Conditional-lethal mutations are those that allow
growth under one condition, but block growth under another condition.
The most common type of conditional-lethal mutation used is a
temperature-sensitive mutation. A temperature-sensitive mutant can grow at one
temperature, the permissive temperature, but it cannot grow at another, the
nonpermissive temperature. In general, the term temperature-sensitive mutation
refers to mutations that impair growth at an elevated temperature. This class of
mutation might also be referred to as heat-sensitive mutations.
In addition, one can also sometimes identify cold-sensitive mutations. The
temperatures used for these classes of mutations might vary, depending on the gene
being studied. In general, for S. cerevisiae, a permissive temperature is likely to be
30°C, and nonpermissive temperatures might be 37°C and 15°C.
In addition to random mutagenesis, sometimes investigators will want to make
specific mutations to test a predicted role or activity of a gene product. For some
genes, the sequence of the gene will suggest that the gene product possesses a
particular biochemical activity. For example, amino acid sequence motifs have been
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identified that confer protein kinase activity upon the protein. In genetic studies of
such a gene, specific mutations that were predicted to affect the protein in a
particular fashion (such as affecting ATP binding) would be of interest. Such
mutations can serve to test particular hypotheses of the role of protein motifs.
Isolating mutations by gene-specific mutagenesis requires several steps. The first
step, the mutagenesis itself, is performed on the isolated gene, usually in the form of
a plasmid. For random mutagenesis, different types of chemical mutagens can be
used, such as hydroxylamine or nitrous acid. An alternative and convenient method
for random mutagenesis of a gene is by the misincorporation of DNA bases, using an
error-prone polymerase chain reaction method. This method relies on the high
frequency of homologous recombination in S. cerevisiae. Site-specific mutations can
be constructed by several standard procedures, usually involving specific
oligonucleotides that contain the mutation of interest combined with PCR.
After mutagenesis, the mutagenized plasmids are used to transform S. cerevisiae.
The transformants are then screened in vivo for mutant phenotypes. In a case where a
gene is essential for viability, a special procedure enables investigators to make
strains in which the only copy of the gene is the potentially mutant one.
Pictured in Figure 7.22, this method is called a plasmid shuffle. For a plasmid
shuffle, the S. cerevisiae strain that is used to screen for the new mutations has the
gene of interest deleted. To maintain viability, it also initially contains a wild-type
copy of the gene on an autonomous plasmid. The plasmid has the S. cerevisiae URA3
gene as its selectable marker. This particular marker is important because one can
select against the wild-type gene by using the compound 5-fluoroorotic acid (5FOA),
which is toxic to any cells that contain the URA3 gene product. Because the plasmid
is lost at a detectable frequency, we can grow the transformants on a medium that
contains the 5FOA and easily identify cells that have lost the wild-type gene. After
the plasmid shuffle procedure, the strain will have only the mutagenized plasmid,
allowing us to screen for mutant phenotypes.
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Figure 7.22 The plasmid shuffle is a method to mutagenize a
cloned, essential gene and to identify conditional mutations.
For example, if we were screening for temperature-sensitive mutations, the
candidates would be replica plated to screen for mutants that can grow at 30°C but
that fail to grow at 37°C. A plasmid shuffle can be used to screen large numbers of
mutant candidates after random mutagenesis of a gene. It can also be used to test for
possible phenotypes of a site-specific mutation.
The process of random or site-specific mutagenesis of a gene results in the presence
of the mutations on autonomous plasmids. However, investigators will often want to
recombine the mutation into the genome, replacing the wild-type gene. This method,
called gene replacement, allows an exact comparison to be made between a
wild-type S. cerevisiae strain and one that differs only by the mutation being studied.
It is especially useful in cases where one cannot select for the mutant allele (unlike
the case in Figure 7.21).
One method for gene replacement involves two recombination steps, shown in
Figure 7.23. In a two-step gene replacement, the mutation of interest is constructed
in an integrating (YIp) plasmid that contains the yeast URA3 gene as a selectable
marker. The DNA is used to transform a ura3 mutant under conditions where the
plasmid will integrate at the gene of interest, resulting in the structure diagrammed.
Because the gene of interest is now duplicated, the plasmid can be excised by
homologous recombination. If the crossover occurs in the correct position, the
remaining allele of the gene will be the mutant allele, resulting in an exact gene
replacement.
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Figure 7.23 The two-step gene
replacement is used to replace a
wild-type gene with any mutant allele
constructed in vitro.
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References
2866. Giaever et al. (2002). Functional profiling of the S. cerevisiae genome. Nature 418, 387-391.
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YEAST GENETICS
Fred Winston
7.19 Protein identification can lead directly to
genetic analysis in S. cerevisiae
Key Terms
An epitope tag is a short amino acid sequence that is recognized by a commercially
available antibody. When fused to a protein of interest, an epitope tag allows
identification of the protein by the available antibody.
Key Concepts
• Identifying an S. cerevisiae protein allows direct identification of the gene that
encodes that protein, which in turn allows genetic analysis to be performed.
In addition to using the traditional methods of mutant screens and selections to
identify genes of interest, we can also begin to identify them by using biochemistry.
That is, we can purify a protein of interest and then, because the complete genome
sequence of S. cerevisiae is known, we can identify the gene that encodes that
specific protein. Following gene identification, genetic analysis can be performed to
analyze the role of the protein in vivo.
Traditionally, biochemical identification of a protein has been based on the
investigator's having an assay for a particular biochemical activity. For example, we
might have an assay for an enzymatic step in amino acid biosynthesis, or for the
occurrence of a regulatory function, such as the binding of a protein to a particular
DNA sequence. The existence of an assay allows purification of a protein by
classical fractionation methods, based on its charge, size, and shape.
Sometimes a biochemical activity relies on a single protein. In other cases, the
activity depends on a complex of proteins, such as for RNA polymerase. In this
situation, the fractionation methods must be able to preserve the association of the
proteins required for the activity.
A second method to purify proteins, used more recently, does not rely on having an
activity assay available. In this case, we can try to identify a protein complex by
beginning with the knowledge of at least one of the genes that encodes a member of
the putative complex. Because proteins often act in complexes, this is a frequently
used approach. The gene being studied is often one found by the type of mutant
analysis described in earlier sections.
We can purify the complex by first fusing the gene of interest to a DNA sequence
encoding an epitope tag. An epitope tag is a short amino acid sequence, generally
eight to eleven amino acids long, that is recognized by a commercially available
antibody. Thus, a protein that contains an epitope tag can be identified by using the
antibody to the tag. The gene fusion is constructed using PCR and yeast
transformation, in a method similar to the one described earlier in this chapter for
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constructing deletion mutations.
One important consideration is whether the gene maintains its wild-type function
after fusing to the epitope-tag sequence. This can depend on the particular epitope
tag, its position in the gene (usually either at the 5′ or 3′ end of the gene), and the
gene of interest. We only want to analyze cases where wild-type function is
maintained; otherwise, the subsequent studies may analyze a mutant situation and be
misleading. Following the fusion, standard methods should be used for making
extracts of yeast cells and purifying the protein that contains the epitope tag. Usually
this is done using a resin coupled to the antibody that recognizes the epitope tag. If
we use conditions that preserve protein-protein interactions, we can often purify the
entire complex in which the epitope-tagged protein exists. After purifying the
complex, we can determine the number and approximate molecular weights of the
proteins by using polyacrylamide gel electrophoresis. Figure 7.24 depicts this
process.
Figure 7.24 DNA encoding an epitope tag is fused to the YFG1 gene. Yeast cells
are then grown up and gently broken open. The Yfg1 protein and its interacting
partners are purified using the epitope tag and an antibody that recognizes the tag.
The identity of the interacting proteins is determined by mass spectrometry.
After a protein is purified by one of these methods, the gene that encodes it can be
identified using two types of information:
• Part of the amino acid sequence of the protein, or
• The determination of the masses of peptides derived from the protein.
We can determine amino acid sequences at the amino-terminal ends of proteins, for
lengths up to about twenty amino acids. Alternatively, it is possible to cleave a
protein into peptides with a specific protease, such as trypsin, and then to ascertain
the precise mass of each peptide by a technique called mass spectrometry. Either of
these methods provides a characterization of the protein that will be specific for that
one protein among all encoded by the yeast genome. Because the predicted protein
product of each gene of S. cerevisiae is known, this allows us to identify the gene
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encoding the protein. This type of approach, which takes advantage of the genome
sequence, has been used on a large scale for S. cerevisiae (2260).
After identifying the gene(s) encoding the protein(s) of interest, we can do several
types of genetic analyses. For example, if a protein was purified based on its ability
to bind to the regulatory region of a particular gene (the target gene), we can make a
mutation in the gene encoding the DNA-binding protein. This will allow us to see if
it alters expression of the target gene. Alternatively, if a group of proteins was
identified as a complex, we can establish whether a mutation in each of the genes
causes the same mutant phenotype. In these ways and others, one can test for the in
vivo roles of proteins after their biochemical identification in vitro. The purification
of a protein complex and the subsequent identification of its members is one way to
identify relevant protein-protein interactions in vivo. (For other methods to identify
protein-protein interactions, see next section.)
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References
2260. Gavin, A. C., et al. (2002). Functional organization of the yeast proteome by systematic analysis
of protein complexes. Nature 415, 141-147.
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YEAST GENETICS
Fred Winston
7.20 Several methods exist to identify genetic
interactions among genes in yeast
Key Concepts
• Often, genetics can identify interactions, including those that happen infrequently
or between rare functions.
• Several methods exist to find such interactions.
After a yeast gene has been identified as playing an important role, it is of great
interest to identify other genes with which it might interact. The genes may encode
related functions or physically interacting proteins. Yeast genetics has developed
several methods that are commonly employed for this goal. The purification and
identification of members of a protein complex were described in an earlier section.
That approach, if successful, will identify proteins that are physically associated.
However, not all protein-protein interactions are strong enough for biochemical
approaches to identify them. Furthermore, some proteins may interact functionally,
but not physically. For example, two different proteins may regulate the same set of
genes, yet they may not physically interact with each other. To identify interactions
that would be difficult or impossible to find biochemically, there are many methods
available to yeast geneticists. Such methods include isolation of suppressors,
identification of synthetic lethal interactions, and two-hybrid analysis.
Last updated on January 27, 2004
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YEAST GENETICS
Fred Winston
7.21 Suppressor analysis is a proven method to
identify interacting genes
Key Terms
Missense mutations change a single codon so as to cause the replacement of one
amino acid by another in a protein sequence.
A high-copy-number suppressor allows a mutation in one gene to be suppressed by
overexpression of a second gene that is present on a high-copy-number plasmid.
A classic approach to identifying interacting genes is the isolation and analysis of
extragenic suppressors. An extragenic suppressor is a mutation in a gene that is
distinct from the gene with the initial mutation. Extragenic suppressors of an
interesting mutation are identified in two general steps.
The first step is the isolation of "revertants" of the original mutant—that is, strains
that no longer have the original mutant phenotype. We can isolate such revertants by
either a screen or a selection. A revertant strain could arise by either true reversion of
the mutation or by a second mutation in an interacting gene, an extragenic
suppressor, that compensates for the defect of the original mutation. One example of
this type of extragenic suppressor was given earlier when we described the epistasis
tests. In that case, a mutation in one gene suppressed the defect caused by a mutation
in another gene.
True revertants and extragenic suppressors are distinguished by the second step. This
is a genetic cross of the "revertant" by a wild-type strain, followed by tetrad analysis,
shown in Figure 7.25. For true revertants, all tetrads will be PD tetrads, since all
spores will inherit a wild-type allele. In contrast, for extragenic suppressors, a
different pattern will be seen because there will almost certainly be recombination
between the suppressor mutation and the original mutation. Therefore, some spores
will inherit only the original mutation. If the original and suppressor mutations are
unlinked, the ratio will be the expected 1:1:4 for PD:NPD:TT. The analysis of
suppressor mutations has been an extremely powerful tool in genetic analysis of
yeast, as well as of other organisms, particularly prokaryotes. (For two good reviews
of suppressor analysis in yeast see 2909 and 2920.)
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Figure 7.25 A true revertant will be completely linked to the initial mutation,
producing all PD tetrads. An extragenic suppressor mutation will segregate
from the initial mutation, resulting in PD, NPD, and TT tetrads.
The nature of the suppressor mutation will often depend on the type of mutation that
is being suppressed. As Figure 7.26 shows, a missense mutation that alters the
conformation of a protein might be suppressed by a compensating change in an
interacting protein. In this case, then, the suppressor mutation identifies an
interacting gene product, often the goal of suppressor analysis. In contrast, a deletion
mutation that removes the coding region of a gene cannot be suppressed by a change
in an interacting protein. Only a "bypass" suppressor that compensates for the
complete loss of the initial gene product can suppress a deletion mutation.
Informational suppressors will often suppress nonsense mutations. We must take all
of this into consideration when planning a project to study a gene by suppressor
analysis, and must carefully consider the molecular nature of the initial mutation.
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Figure
7.26
Suppression
between
interacting proteins. In the example shown,
two proteins, A and B, normally interact. In
the a mutant, this interaction is impaired and
there is a mutant phenotype. However, a
suppressor mutation that alters B restores the
interaction with a, resulting in a wild-type
phenotype.
In addition to extragenic suppressors, a second type of suppressor that we can screen
for is called a high-copy-number suppressor. This screen identifies genes that,
when overexpressed, suppress the defect caused by the mutation in the initial gene.
For example, imagine that a mutation weakens an interaction between two proteins
because it reduces the affinity between the mutant protein and the second protein.
This results in a mutant phenotype. If the level of the second protein is now
increased, it might overcome the reduced affinity, resulting in a wild-type phenotype.
High-copy-number suppression can occur by other mechanisms as well. Screens for
high-copy-number suppressors are done by using an S. cerevisiae genomic library in
a high-copy-number (2-micron) vector to transform the mutant of interest. The
transformants are then screened for those with a wild-type phenotype. The plasmids
in these candidates are isolated and tested to see if they contain the gene
corresponding to the mutant gene (not the desired class) or to a different gene that
might be a high-copy-number suppressor. A gene in the latter category would likely
encode a protein that would interact with the mutant protein.
Genes identified by suppressor analysis can be cloned and studied using all the types
of genetic, molecular, and biochemical approaches described in this chapter. Often,
the study of a gene identified as a suppressor will shed light not only on the
suppressor but on the gene containing the initial mutation. Suppressor analysis is a
powerful tool for yeast geneticists.
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2909. Forsburg, S. L. (2001). The art and design of genetic screens: Yeast. Nat. Rev. Genet. 2, 659-668.
2920. Prelich, G. (1999). Suppression mechanisms: themes from variations. Trends Genet. 15, 261-266.
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7.22 Synthetic lethal interactions can identify genes
with related functions
In addition to the isolation of suppressor mutations, a second genetic approach to
identify interacting genes is called a synthetic lethal screen. This type of screen
identifies pairs of mutations in which both are required to cause inviability. The
rationale for identifying and studying such genes is that if each of two mutations
impairs an aspect of an essential process, the double mutant should have a more
severe phenotype than either of the single mutants. This double-mutant lethality is
called synthetic lethality. Thus, a synthetic lethal screen is the opposite of a
suppressor screen, in which the second mutation causes a wild-type phenotype.
A synthetic lethal screen begins with an S. cerevisiae strain that contains a mutation
in the gene of interest, and also a plasmid that has both a wild-type copy of that gene
and the URA3 gene as a selectable marker. Figure 7.27 illustrates this. By
performing mutagenesis and screening mutagenized colonies that have lost the
plasmid, we can identify second mutations that cause inviability when the wild-type
copy of the gene on the plasmid is missing.
Figure 7.27 A synthetic lethal screen identifies functional interactions in an
essential process.
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Synthetic lethal screens can identify genes that are related in different ways, as
Figure 7.28 and Figure 7.29 demonstrate. Three possible relations are those among
genes in a regulatory pathway, genes whose products interact together in a complex,
and genes whose products independently contribute to the same process.
Figure 7.28 Two types of interactions are diagrammed. In
one possibility, two genes, TUB1 and TUB3, each encodes a
version of the essential protein alpha-tubulin, with TUB1
providing most of the product. Some point mutations in
TUB1 that destabilize the interactions of resulting protein are
not lethal. However, in combination with a tub3 null allele,
there is not enough functional alpha-tubulin and the result is
lethality.
Figure 7.29 Several proteins interact in a complex that carries out an
essential function. A mutation in X, Y, or Z reduces the activity of
this complex, but still allows enough for viability. However, a double
mutant that impairs two proteins reduces the activity below a
threshold required for viability, resulting in synthetic lethality.
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Fred Winston
7.23 Two-hybrid analysis is a powerful way to
identify protein-protein interactions in vivo
Another popular approach for detecting interactions is a two-hybrid analysis (952).
This method is designed to detect physical interactions between two proteins in vivo.
Two-hybrid analysis relies on two characteristics of transcriptional activator proteins:
• Many transcriptional activator proteins contain two functional domains: a
DNA-binding domain and a transcriptional activation domain.
• These two domains can often be physically separated. That is, if the
DNA-binding domain is present on one protein and the transcriptional activation
domain is present on a second, interacting protein, interaction of the two proteins
will activate transcription.
Thus, we can make transcriptional activation dependent on a particular
protein-protein interaction in vivo.
On the basis of these properties, we can screen S. cerevisiae for all proteins that
physically interact in vivo with a protein of interest. This is illustrated in Figure 7.30.
First, recombinant DNA methods are used to create a plasmid encoding a hybrid
gene that expresses the protein of interest fused to a known DNA-binding domain.
The most commonly used DNA-binding domains are both well characterized: one is
from the yeast transcriptional activator Gal4 and the other from the E. coli repressor
LexA.
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Figure 7.30 Proteins that interact with each other can be identified by
their ability to activate transcription by linking a DNA-activating
domain to a DNA-binding domain.
A recombinant library is then made in which random S. cerevisiae DNA fragments
are cloned into a vector where they can potentially be fused to a well-characterized
transcriptional activation domain. Often, the Gal4 transcriptional activation domain
is used in this type of vector.
Next, an S. cerevisiae strain is used in which the binding sites for the DNA-binding
domain (for example, Gal4 binding sites) are placed 5′ of a reporter gene. To perform
the two-hybrid screen, the plasmid that encodes the DNA-binding domain fusion
protein is first used to transform S. cerevisiae. Then, the library of fusions to the
activation domain is used to transform the same strain. Individual transformants are
screened or selected (depending on the reporter used) to identify those in which
transcription of the reporter has been activated. Such cotransformants are candidates
to have a two-hybrid interaction. The determination of the DNA sequence in the
activator plasmid identifies the putative interactor. Subsequent biochemical and
genetic tests can confirm a physical interaction.
Besides suppressor analysis and two-hybrid analysis, yeast geneticists employ many
other types of approaches to identify functionally related genes. Which method is the
best one to use? To answer that question requires being able to predict the future. We
never know what genes will be discovered; however, the long and successful history
of these and related approaches holds the promise of new and exciting insights.
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References
952. Fields, S. and Song, O. (1989). A novel genetic system to detect protein-protein
interactions. Nature 340, 245-246.
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7.24 Functional genomics studies of S. cerevisiae
Key Terms
Whole-genome expression analysis is a method in which the mRNA level of every
gene is determined.
Bioinformatics describes computational methods to study the data from
whole-genome expression analysis.
Key Concepts
• The knowledge of the full genomic sequence of S. cerevisiae enables us to perform
several types of genomewide analyses.
The information in the complete sequence of an organism's genome allows us to
apply methods that analyze the entire genome. This is particularly true of
transcription. As the first eukaryote to have its genome completely sequenced, S.
cerevisiae has provided a model system for functional genomic studies. These
studies have illustrated powerful methods to analyze mutations that affect gene
expression.
One of the most attractive functional genomic methods is whole-genome expression
analysis(2717). Using these means, we can analyze the level of the mRNA for every
gene in S. cerevisiae. By comparing mRNA levels between a wild-type strain and a
mutant, or between different mutants, we can discern the full extent of a mutant
phenotype at the level of mRNA expression.
As Figure 7.31 shows, the general method for whole-genome expression analysis
entails using microarrays. In this technique, the DNA sequence that corresponds to
each S. cerevisiae gene is synthesized and then fixed in an ordered pattern on a glass
slide. Following this is the preparation of mRNA populations from the two S.
cerevisiae strains (wild type and mutant) that will be compared. Each mRNA
population is used to make fluorescently labeled cDNA so that the levels of cDNA
and mRNA will correspond. The wild-type and the mutant cDNA are given different
fluorescent labels. The two pools of cDNA are then mixed and hybridized to the
microarray and processed for analysis. The amount of hybridization of each
population to each spot in the microarray is proportional to the level of the mRNA in
vivo from each of the two strains. When two strains such as a wild type and a mutant
are compared, the relative levels are expressed as a ratio. This reveals any change in
the levels of mRNA. While this type of approach is still relatively new, several
studies have already been done. These include one that has identified all of the S.
cerevisiae genes that are specifically expressed during meiosis (2912).
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Figure 7.31 Whole-genome expression analysis compares the
mRNA population of a mutant strain to wild type. Microarray
image kindly provided by Tom Volkert, Whitehead Institute, Center
for MicroArray Technology.
The analysis of microarray data has given birth to a new field of computational data
analysis. Dubbed bioinformatics, this has rapidly grown into a major field of
research in modern biology. One of the goals of bioinformatics is to make
predictions based on computational analysis of microarray data. For example,
analyzing a set of genes that are coregulated might identify common DNA sequences
in the genes' regulatory regions. These might serve as cis-acting regulatory elements.
This hypothesis could then be tested experimentally.
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References
2717. Lashkari, D. A., DeRisi, J. L., McCusker, J. H., Namath, A. F., Gentile, C., Hwang, S. Y., Brown,
P. O., and Davis, R. W. (1997). Yeast microarrays for genome wide parallel genetic and gene
expression analysis. Proc. Natl. Acad. Sci. USA 94, 13057-13062.
2912. Chu, S., DeRisi, J., Eisen, M., Mulholland, J., Botstein, D., Brown, P. O., and Herskowitz,
I. (1998). The transcriptional program of sporulation in budding yeast. Science 282, 699-705.
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7.25 S. cerevisiae and S. pombe have been
invaluable organisms for elucidating cell cycle
control
Key Concepts
• Studies of cell cycle control in yeast have led the way to understanding this process
in all eukaryotes.
Studies of cell cycle control in two yeasts, S. cerevisiae and S. pombe, have greatly
enhanced our understanding of cell division in all eukaryotes. This success is based
on the ability to isolate and analyze mutants that are blocked in specific steps of cell
division. The significance of this work is highlighted by the award of the 2002 Nobel
prize in medicine or physiology to Leland Hartwell and Paul Nurse, the pioneers in
the field of cell cycle control studies for S. cerevisiae and S. pombe, respectively.
Beginning in 1969, Leland Hartwell and colleagues took a novel approach to
studying a large collection of temperature-sensitive mutants of S.
cerevisiae (2038)(824) (See also Great Experiments 4 Cell cycle genes). These
mutants had been isolated solely on the basis of their ability to grow at a normal
temperature (30°C), but not at a high temperature (36°C). This limitation indicated
that they had temperature-sensitive mutations in an essential gene.
When the mutants were examined in the microscope after being shifted to a
nonpermissive temperature, it was discovered that some of them had arrested growth
with a uniform cell morphology. That is, they exhibited a particular bud size, or no
bud. For any given type of mutant, most cells had arrested at the same position in the
cell cycle, indicating that the mutant gene product was normally necessary at that
point in the cycle. One class of mutants arrested as unbudded cells, another class
with small buds, still another with large buds, and so on. All of the mutants that
behaved in this fashion were named cell division cycle (cdc) mutants.
The ability to isolate cdc mutants in S. cerevisiae led to the hypothesis that specific
sets of genes were required to pass through sequential stages of the cell cycle. For
example, a cdc mutation that caused cells to arrest in an unbudded form would
suggest that that specific gene was needed at the beginning of the cell cycle. A
mutation in a gene required for chromosome segregation might result in a large,
budded cell. This would indicate that the cell was blocked after DNA replication.
This type of analysis of a large number of mutants led to the understanding that many
gene products are required for passage through many different points in the cell
cycle.
Genetic analysis of numerous cdc mutants showed that cell division in S. cerevisiae
consisted of two major pathways, one important for nuclear division and the other for
cell budding and cytokinesis. In this analysis, cdc mutants were examined for their
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morphology, DNA replication, and nuclear division.
Genetic analysis of cdc mutants also demonstrated that these are dependent
pathways; that is, later events do not happen in the absence of earlier events. For
example, mitosis does not occur if DNA replication has not taken place. This large
body of work, which has involved many S. cerevisiae labs, has provided the
foundation for studies of all eukaryotic cell division for over thirty years.
The relevance of the studies of yeast cdc mutants became evident when biologists
perceived the strong conservation throughout eukaryotes of many types of cell
functions. One particularly important breakthrough involved studies of the yeast S.
pombe, which is distantly related to S. cerevisiae. Cell cycle studies of this yeast
identified the cdc2 gene as being essential for the initiation of the cell cycle (2872).
In a successful attempt to identify the human gene that serves a function equivalent
to that of S. pombe cdc2, Paul Nurse and his colleagues cloned the human gene by
complementation of an S. pombe cdc2 temperature-sensitive mutant (2041) (See also
Great Experiments 2 The discovery of cdc2 as the key regulator of the cell cycle).
Subsequent work demonstrated that human cdc2 encodes a biochemical activity that
is essential for cell division.
One major concept to emerge from studies of the S. cerevisiae cell cycle is that of
checkpoints that control the dependency of later events on the successful completion
of earlier events (319). In principle, either a set of controls or the appearance of
appropriate intermediate steps could regulate the dependency of events during the
cell cycle.
In 1988, Ted Weinert and Leland Hartwell demonstrated that a set of controls
existed. In these studies, they screened for mutants that would allow the cell cycle to
progress under conditions in which it is was normally inhibited by DNA damage
(843). They identified a mutation in the RAD9 gene that allowed such an aberrant
progression.
Since this landmark discovery, evidence for several other types of cell cycle
checkpoints has been discovered, not only in S. cerevisiae but throughout eukaryotes.
Studies of the cell cycle in both S. cerevisiae and S. pombe were of critical
importance in clarifying the process of cell division in eukaryotes. Furthermore,
these studies have helped to elucidate cellular processes that have gone awry in
cancer cells (2913).
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Reviews
319. Hartwell, L. H. and Weinert, T. A. (1989). Checkpoints: controls that ensure the order of cell
cycle events. Science 246, 629-634.
2913. Hartwell, L. H., and Kastan, M. B. (1994). Cell cycle control and cancer. Science 266,
1821-1828.
References
824. Hartwell, L., Culotti, J., Pringle, J. R., and Reid, B. J. (1974). Genetic control of the cell division
cycle in yeast. Science 183, 46-51.
843. Weinert, T. A., and Hartwell, L. H. (1988). The RAD9 gene controls the cell cycle response to
DNA damage in S. cerevisiae. Science 241, 317-322.
2038. Hartwell, L. H., Culotti, J., and Reid, B. (1970). Genetic control of the cell-division cycle in yeast.
I. Detection of mutants. Proc. Natl. Acad. Sci. USA 66, 352-359.
2041. Lee, M. G. and Nurse, P. (1987). Complementation used to clone a human homologue of the
fission yeast cell cycle control gene cdc2. Nature 327, 31-35.
2872. Nurse, P., Masui, Y., and Hartwell, L. (1998). Understanding the cell cycle. Nat. Med. 4,
1103-1106.
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7.26 The isolation of mutants unable to mate
elucidated the mating pathways in yeast
Key Terms
Sterile mutants are yeast mutants that cannot mate.
Key Concepts
• The study of mating type control in S. cerevisiae has illuminated the understanding
of a signaling pathway.
One of the best-understood cases of gene regulation in S. cerevisiae is the
determination of mating type. Recall that haploid S. cerevisiae strains exist in one of
two mating types, either a or α. The genetic locus that determines the a or α mating
type, the MAT locus, has two possible states: MATa or MATα. Each locus contains
two genes that confer a or α cell identity. In S. cerevisiae, the major aspects of
mating type control—that is, how a cell becomes an a or α type—and the steps
required for mating were determined by mutant analysis.
In steps during mating described earlier in Figure 7.4, each requires the regulated
expression of several genes (see Genetics 7.5 The S. cerevisiae life cycle). In 1974,
two S. cerevisiae geneticists, Vivian Mackay and Thomas Manney, set out to identify
the genes required for the yeast to mate by isolating mutants that were unable to do
so. These were called sterile mutants, and the genes identified were named STE
genes (2870)(2914). Mackay and Manney isolated many such mutants. Using
methods similar to those described earlier in this chapter, they placed the mutants
into complementation and linkage groups.
Because both complementation and linkage analysis in S. cerevisiae require diploids
and hence mating, how was this accomplished for ste mutants? Some of the ste
mutants were conditional, meaning they were sterile at one temperature but able to
mate at another. Other mutants were leaky, that is, they still had some function, but it
was greatly reduced. Such mutants were able to mate at low frequency, allowing
diploid formation.
From their analysis, Mackay and Manney identified three general classes of ste
mutants:
• Those that caused sterility in either a or α cells;
• Those that caused sterility only in a cells; and
• Those that caused sterility only in α cells.
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This classification was possible because all of the mutants were still able to mate,
although at a very low frequency. Therefore, Mackay and Manney were able to
examine the phenotype of each mutation in both MATa and MATα genetic
backgrounds and test its effect on mating. In addition to the finding of three classes
of mutants, their work resulted in two other significant discoveries. First, their
research demonstrated that many genes are important for normal mating, not just the
previously identified MAT locus. Second, while they isolated ste mutations at the
MATα locus, they were unable to identify any ste mutations at the MATa locus. This
suggested that the two different loci did not contribute equivalent functions within a
similar regulatory network. Later work showed that this hypothesis was correct.
Many of the STE genes required for mating in either mating type have been shown to
be involved in a signal transduction pathway. This pathway is initiated by the
binding of a mating pheromone to its receptor—for example, the binding of an α
factor (produced by MATα cells) to an α-factor receptor (on the surface of MATa
cells). This binding triggers a series of steps, via protein kinases. These steps lead to
two cellular responses: cell cycle arrest and the induction of gene transcription
required for the mating process. This signal transduction pathway is described in
greater detail in Molecular Biology.
The study of cell type in S. cerevisiae has been a remarkably rich area of
investigation. In addition to the signal transduction pathway described in this section,
great advances were made in transcriptional control, gene silencing, and directed
recombination to switch mating type. These are all described in detail in Molecular
Biology 18 Rearrangement of DNA.
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References
2870. Mackay, V., and Manney, T.R. (1974). Mutations affecting sexual conjugation and related
processes in S. cerevisiae. I. Isolation and phenotypic characterization of nonmating
mutants. Genetics 76, 255-271.
2914. MacKay, V., and Manney, T.R. (1974). Mutations Affecting Sexual Conjugation and Related
Processes in S. cerevisiae. II. Genetic Analysis of Nonmating Mutants. Genetics 76, 273-288.
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7.27 Mitochondrial genetics in S. cerevisiae
Key Concepts
• S. cerevisiae that lack mitochondrial DNA are viable.
• Segregation of mitochondrial DNA mutations is non-Mendelian.
Mitochondria are organelles that are the sites of respiration and oxidative
phosphorylation. A mitochondrion contains its own genome and uses its own
transcription and translation machinery. In fact, mitochondria can use a genetic code
that is slightly different from the genetic code of nuclear genes (see Molecular
Biology 7.7 There are sporadic alterations of the universal code).
Mutations in S. cerevisiae mitochondrial DNA (mtDNA), including complete loss of
mtDNA, cause defects in respiration but allow viability. Extensive genetic analysis
of S. cerevisiae mutations that alter mitochondrial function has helped to elucidate
that function, and has also established the inheritance patterns of different classes of
mitochondrial mutants. In addition, the mechanism by which proteins from the
cytoplasm are directed to the mitochondria and the way the mitochondria control
their gene expression have both been studied extensively.
S. cerevisiae has served as an ideal organism for studying mitochondrial function
because members of that species that have completely lost mtDNA are still viable as
long as they have a carbon source, such as glucose, that allows fermentation. (Note
that such mutants still form mitochondria, even though these mitochondria do not
contain any mtDNA.) S. cerevisiae mutants that have lost respiration function fail to
grow on carbon sources that require respiration, such as lactate or ethanol. This
means that mutations that impair mitochondrial function can be isolated and
identified by their conditional growth, depending on the particular carbon source.
The mitochondrial DNA of S. cerevisiae is 75 kilobases in length, approximately five
times longer than human mtDNA. S. cerevisiae mtDNA contains genes required for
some mitochondrial functions. These genes encode tRNAs, rRNAs, cytochrome
oxidase, and an ATPase. The remaining mitochondrial components are encoded in
nuclear genes, including the genes encoding the subunits for mitochondrial RNA
polymerase and mitochondrial ribosomes.
Because both mitochondrial and nuclear genes are necessary for mitochondrial
function in S. cerevisiae, mutations in either mtDNA or nuclear DNA can impair that
function. Each class of mutation behaves in a distinct fashion genetically, indicating
cytoplasmic or nuclear inheritance. The nuclear genes required for mitochondrial
function are called PET genes. This stands for petite, the general term for S.
cerevisiae mutants that are defective for mitochondrial function. Mutations in PET
genes segregate 2:2, as expected for any single nuclear mutation.
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Mutations in mtDNA, called rho mutations, segregate in a different fashion from
nuclear pet mutations, indicating cytoplasmic inheritance. Furthermore, different
classes of rho mutations exhibit distinct segregation patterns. First, let us consider
rho mutations that are the result of the complete loss of mtDNA. These are called
rho0 mutations. If a rho0 strain is crossed by a wild-type strain, the cytoplasms of the
two parents become mixed in the diploid. As you can see in Figure 7.32, after
sporulation, wild-type mitochondria are transmitted to all four spores of each tetrad.
Therefore, the segregation is 4:0 rho +:rho0. For most point mutations in mtDNA,
called rho – mutations, the same type of cytoplasmic inheritance is evident.
Figure 7.32 In this cross, one parent is wild
type (rho+) and the second parent lacks
mitochondrial DNA (rho0). When the strains
are crossed, all spores in a tetrad inherit
mitochondria. Cytoplasmic inheritance is
indicated by the 4:0 segregation pattern.
Another class of rho – mutations arises from a deletion of most of the mtDNA. For
reasons that are not yet clear, the remaining mtDNA amplifies so that its total size is
equal to that of wild-type mtDNA. In many cases, this class of rho – mutants displays
a different segregation pattern. By an unknown mechanism that may have to do with
the rate of DNA replication, the mutant rho – genome can outcompete the wild-type
mtDNA. This results in many more mutant mitochondria than wild-type
mitochondria, so that most or all of the spores inherit the mutant rather than the
wild-type form. The reason for this may be faster replication of the mutant
mitochondrial genome. Some rho –mutants of this class have only a partial effect,
producing a mixture of wild-type and mutant progeny. The relative frequency of the
two classes is called the degree of suppressiveness.
Studies of both nuclear and mitochondrial genes have benefited greatly from the
ability to transform them. Transforming S. cerevisiae mitochondria required a special
technique, as the usual method for transformation in that species did not work. In
1988, the transformation of S. cerevisiae mitochondria was achieved by using a
"biolistic," or biological ballistic (2718). This technique entails high-velocity
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microprojectile bombardment, using very small particles (1 µm in diameter) coated
with the transforming DNA. A "gun" fires the particles at a speed sufficient to enter
the cells and disrupt the mitochondrial membrane to allow delivery of the DNA. It is
not, however, so violent as to kill the S. cerevisiae cells. Using this method, rho0
strains can be transformed with specific constructs, allowing their analysis in vivo.
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References
2718. Butow, R. A., Henke, R. M., Moran, J. V., Belcher, S. M., and Perlman, P.
S. (1996). Transformation of S. cerevisiae mitochondria using the biolistic gun. Methods
Enzymol. 264, 265-278.
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7.28 Relevance of yeast studies to human health
Key Concepts
• Many human genes associated with disease have yeast counterparts.
• Studies of gene function in S. cerevisiae can illuminate the factors important for
virulence in the pathogenic yeast C. albicans.
Over the past several years, it has become clear that studies in yeast can be directly
relevant to understanding human biology, and are thus relevant to human health. In
this section, we will cover two areas that demonstrate these attributes.
Over the past twenty years, it has become obvious that there is a remarkable degree
of conservation among all eukaryotes in fundamental aspects of growth. We have
already described how this is true for the cell cycle. In other processes, too, including
the translation of RNA and the transcription, replication, and repair of DNA,
conserved factors have been identified in eukaryotes ranging from yeast to humans.
As mutant genes that cause human disease are cloned and sequenced, a significant
percentage of them are found to have homologues in the yeast genome. Such
diseases include colon cancer, cystic fibrosis, Bloom syndrome, and Werner
syndrome. Studying the roles of these gene functions in yeast, where their analysis
can be more rapid and detailed, will help explain the mechanism of the diseases they
cause in humans.
The identification of the first human gene associated with hereditary nonpolyposis
colon cancer (HNPCC) arose from studies of DNA repair in yeast and in bacteria
(2259)(2258). HNPCC had been shown to cause genomic instability in mammalian
cells, suggesting that the disease was caused by a defect in DNA repair. The study of
the E. coli MutS and S. cerevisiae MSH2 genes had shown that they might be related
to this type of DNA repair activity. The human gene hMSH2 was identified as a
homologue of the E. coli MutS and S. cerevisiae MSH2 genes. Studies of this gene in
HNPCC patients identified a mutation and showed that it was associated with
inheritance of the disease gene. Therefore, in this case, the human gene was
identified based on its initial identification and analysis in bacteria and S. cerevisiae.
Its continued study in these microorganisms will aid our understanding of this type
of cancer.
In addition to its use to study homologues of human disease genes, S. cerevisiae can
be used to teach us much more about pathogenic yeasts such as C. albicans. C.
albicans is the most widespread human fungal pathogen, and it can cause
life-threatening infections in immunocompromised patients. Therefore,
understanding more about C. albicans growth and infection could result in important
medical advances.
C. albicans can grow in different states. One of these is similar to that of S.
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cerevisiae, which grows by budding. In addition, C. albicans can grow in
filamentous forms. After it infects mammalian hosts, it can exist in all of these states.
The effect of each one on the organism's virulence in mammalian cells is not well
understood.
One drawback to studying C. albicans is that genetic studies of the species are
extremely difficult. This is because C. albicans grows only as a diploid, and it has no
known sexual cycle. Therefore, most of the attributes of genetic analysis available
for S. cerevisiae are lacking for C. albicans. Mutant screens cannot be performed as
easily as in S. cerevisiae because C. albicans is a diploid. In addition, neither
complementation tests nor linkage tests can be done. Because of these difficulties,
advances in our understanding of C. albicans infections have been very slow.
However, some genetic analysis can be performed in C. albicans because
transformation and gene replacement methods work.
Much of the recent progress in understanding C. albicans' filamentous growth and
pathogenicity is based on studies that originated in S. cerevisiae. Researchers
recently discovered that under certain nutrient limitations, S. cerevisiae can grow in a
filamentous fashion, like C. albicans (2721). Genetic studies have identified several
factors that are required for filamentous growth in S. cerevisiae. Many of the same
factors that are required for haploid S. cerevisiae to respond to mating pheromones
are also required for diploid S. cerevisiae to respond to nitrogen starvation and
undergo filamentous growth.
Many of the genes in S. cerevisiae and in C. albicans are conserved. Thus we can test
whether the same genes that are important for filamentous growth in S. cerevisiae are
also important for filamentous growth in C. albicans. These experiments are
performed by identifying the C. albicans homologue and then constructing null
alleles by gene replacement methods (2722). Because C. albicans can live only as a
diploid, construction of each null mutant requires two copies of the gene to be
deleted. Nevertheless, this task has been accomplished for several C. albicans genes.
The results have identified several genes important for both the species' filamentous
growth and its virulence. Thus, studies in S. cerevisiae have succeeded in moving us
toward a detailed understanding of C. albicans infections.
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References
2258. Fishel, R., Lescoe, M. K., Rao, M. R., Copeland, N. G., Jenkins, N. A., Garber, J., Kane, M., and
Kolodner, R. (1993). The human mutator gene homolog MSH2 and its association with hereditary
nonpolyposis colon cancer. Cell 75, 1027-1038.
2259. Leach, F.S., Nicolaides, N.C., Papadopoulos, N., Liu, B., Jen, J., Parsons, R., Peltomaki, P.,
Sistonen, P., Aaltonen, L.A., Nystrom-Lahti, M., et al. (1993). Mutations of a mutS homolog in
hereditary nonpolyposis colorectal cancer. Cell 75, 1215-1225.
2721. Gimeno, C. J., Ljungdahl, P. O., Styles, C. A., and Fink, G. R. (1992). Unipolar cell divisions in
the yeast S. cerevisiae lead to filamentous growth: regulation by starvation and RAS. Cell 68,
1077-1090.
2722. Liu, H., Kohler, J., and Fink, G. R. (1994). Suppression of hyphal formation in Candida albicans
by mutation of a STE12 homolog. Science 266, 1723-1726.
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YEAST GENETICS
Fred Winston
7.29 What's next?
Studies of yeasts have moved extremely rapidly over the past several years,
accelerated by the S. cerevisiae genome sequence and emerging technologies, such
as microarrays, that allow global analysis of gene expression. Future studies will take
advantage of these new technologies to provide large amounts of data measuring
gene expression, assaying the binding of proteins to DNA, and characterizing all
gene products.
However, even with these technologies, future studies will still require use of the
classical genetic methods that have been the backbone of yeast genetics. For
example, there are still thousands of S. cerevisiae and S. pombe genes whose
functions are completely unknown. While their expression and their gene products
will be characterized by modern methods, it will take mutant analysis and traditional
biochemical analysis to elucidate the actual functions of these genes and their
products.
In addition to progress in studies of S. cerevisiae and S. pombe, studies of yeast
pathogens, including C. albicans and others, will benefit from the new approaches
discussed in this chapter. It is to be hoped that this progress will lead to medical
breakthroughs in understanding fungal infections, while also shedding new light on
how these particular yeasts grow.
Finally, rapid sequencing technologies will allow the genome sequences to be
determined for many different yeasts. Analysis of these sequences will provide
fascinating new insights into the evolution of yeast species.
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YEAST GENETICS
Fred Winston
7.30 Summary
Yeast genetics has been the driving force in the discovery of many fundamental
aspects of eukaryotic cell division and gene expression. This chapter has summarized
classical, molecular, and genomic methods for understanding yeast gene function.
The ability to study yeasts as both stable haploids and diploids, combined with tetrad
analysis, offers powerful tools for genetic analysis. Methods for gene cloning, gene
mutagenesis, and gene replacement allow investigators to carry out detailed
experiments to analyze gene function in vivo. Several other genetic approaches,
including suppressor analysis, synthetic lethal screens, and two-hybrid analysis,
allow the clarification of genetic interactions in vivo. Overall, yeast genetics provides
an ideal system for studies of fundamental eukaryotic processes.
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