DNA Repair
Protocols
Eukaryotic Systems
DNA Repair
Protocols
Eukaryotic Systems
HUMANA PRESS
Methods in Molecular Biology
TM
TM
Methods in Molecular Biology
Edited by
Daryl S. Henderson
VOLUME 113
Edited by
Daryl S. Henderson
HUMANA PRESS
Technical Notes
UV-A, UV-B, and UV-C: This terminology, which divides the ultraviolet
(UV) spectrum into three wave bands, was first proposed in 1932 by the Ameri-
can spectroscopist William Coblentz and his colleagues to begin to address
the problem of standardizing the measurement of UV radiation used in medi-
cine (1,2). Each spectral band was defined “provisionally” and “approximately”
by the absorption characteristics of specific glass filters as follows: UV-A,
400–315 nm; UV-B, 315–280 nm; UV-C, <280 nm (1). Although based on
physical specifications, these definitions were influenced by knowledge of
other UV phenomenology, including biological effects and physical proper-
ties. For example, wavelengths in the UV-B band were known to have potent
erythemic effects, and wavelengths below 290 nm were known to be absent
from sunlight (2) (because they are absorbed by stratospheric ozone). More-
over, the germicidal effects of UV-C wavelengths (principally around 266

nm) from artificial sources were well-recognized (3). Today, the spectral bands
implied by these terms may be found to vary from Coblentz’s original defini-
tions, depending on the discipline. Environmental photobiologists, for example,
generally use the following definitions: UV-A, 400–320, UV-B, 320–290, and
UV-C, 290–200 (4).
Relative centrifugal forces: The g-forces listed in this book are calcu-
lated for the maximum radius unless stated otherwise. For microcentrifuges
similar to Eppendorf’s 5410 and 5415 C models, maximum rotational speed
(14,000 rpm) corresponds to ~12,000g and ~16,000g, respectively.
References
1. Coblentz, W. W. (1932) The Copenhagen meeting of the Second International
Congress on Light. Science 76, 412–415.
2. Coblentz, W. W. (1930) Instruments for measuring ultraviolet radiation and the
unit of dosage in ultraviolet therapy. Br. J. Radiol. 3, 354–363.
3. Gates, F. L. (1930) A study of the bactericidal action of ultra violet light. III. The
absorption of ultra violet light by bacteria. J. Gen Physiol. 14, 31–42.
4. Diffey, B. L. (1991) Solar ultraviolet radiation effects on biological systems. Phys.
Med. Biol. 36, 299–328.
xix
Checkpoint Mutant Screen in
S. pombe
1
1
1
From:
Methods in Molecular Biology, Vol. 113: DNA Repair Protocols: Eukaryotic Systems
Edited by: D. S. Henderson © Humana Press Inc., Totowa, NJ
Isolation of DNA Structure-Dependent
Checkpoint Mutants in
S. pombe

Rui G. Martinho and Antony M. Carr
1. Introduction
Eukaryotic cells have the ability to influence progression through the cell
cycle in response to internal and external inputs of “information”. They do so
by using feedback control mechanisms able to arrest mitosis in response to
different cellular events. Such active mechanisms capable of influencing the
timing of cell-cycle events have been called “checkpoints” (1,2). Cells arrest
progression through the cell cycle if they fail to complete DNA replication or if
their DNA is damaged. The S-phase/mitosis (S-M) checkpoint plays a key role
in the maintenance of the interdependency between S-phase and mitosis. Wild-
type Schizosaccharomyces pombe (fission yeast) cells arrest cell-cycle pro-
gression in response to a DNA replication block, such as that induced by
hydroxyurea (HU), but continue to grow in size, since they are still metaboli-
cally active. These cells are observed to have an elongated phenotype. Mutants
have been isolated in S. pombe that have lost the S-M checkpoint and do not
prevent mitosis if DNA replication during the previous S-phase is incomplete
(3–7). S-M checkpoint mutants do not delay cell-cycle events after exposure to
HU, and will enter mitosis with unreplicated DNA. As a consequence, the elon-
gated phenotype seen for wild-type cells is absent in checkpoint mutants. In-
stead, these mutants show a characteristic “cut” phenotype, where a cell has
entered an abortive mitotic event followed by the formation of a septum through
the nucleus. In these small dead cells, the nucleus is frequently cut in two by
the septum and/or spread unevenly between both daughter cells. S-M check-
point mutants show very low viability in the presence of HU or any other cir-
cumstance that may delay S-phase progression (e.g., in combination with a
thermosensitive DNA replication mutant).
2 Martinho and Carr
Several of these S-M checkpoint mutants are also unable to arrest the cell
cycle in response to DNA damage (6,7). The DNA damage checkpoint arrests
the cell cycle in a dose-dependent manner after exposure to DNA-damaging

agents. It is believed that these delays provide additional time for cells to repair
the damaged genetic material before key transitions are attempted (between
G1 and S-phase, G2 and mitosis, or during DNA replication). Several different
mutants have been isolated in S. pombe that are defective in the DNA damage
checkpoint. In contrast to wild-type cells, which arrest the cell cycle immedi-
ately after DNA damage (and display the previously described elongated phe-
notype), most of these checkpoint mutants do not show any delay in cell-cycle
events after exposure to DNA-damaging agents. One or two of these mutants
are only partially defective (e.g., rad24). As seen for the S-M checkpoint
mutants, the DNA damage checkpoint mutants can also show a cut phenotype
where cells enter unrestrained mitosis with damaged DNA. All these mutants
are highly sensitive to DNA-damaging agents. We refer to these two check-
points (S-M and DNA damage) as DNA structure-dependent checkpoints.
The existence of many genes whose function is required for both checkpoint
controls suggests a significant overlap between these two pathways. The struc-
tural identity between the checkpoint proteins from fission and budding yeast
suggests that these pathways have analogs in mammalian cells. This is sup-
ported by the growing number of human genes found to be homologous to
yeast checkpoint genes (8).
We describe below methods for:
1. Generating mutants of S. pombe.
2. Screening those mutants for putative DNA structure-dependent checkpoint defects.
3. Distinguishing between S-M and DNA damage checkpoint deficiencies.
4. Further characterizing checkpoint mutants.
2. Materials
2.1. Media
1. Yeast extract medium (YE): 5 g/L Difco (Detroit, MI) yeast extract, 30 g/L glu-
cose, supplemented as required with 100 mg/L leucine, adenine, lysine, uracil,
and histidine.
2. Yeast extract agar medium (YEA): YE plus 20 g/L Difco agar.

3. YEP: YE plus 20 g/L Difco Bacto-peptone.
4. Phloxin B agar (YEA + P): YEA plus 0.02 mg/mL Phloxin B (Sigma, Dorset,
UK). Phloxin B is stored as a stock solution at 20 mg/mL. It should be added after
the medium is autoclaved and cooled.
5. YEA + P with HU: YEA + P containing 10 mM HU. HU is kept as a 1 M stock
solution stored at –20°C. It is filter-sterilized and added to autoclaved, cooled
medium.
Checkpoint Mutant Screen in
S. pombe
3
2.2. Additional Reagents and Equipment
1. Ethyl methanesulfonate (EMS is listed in Sigma’s catalog as methanesulfonic
acid ethyl ester).
2. 254-nm UV source (e.g., germicidal lamp).
3. 4,6-Diamino-2-phenylindole (DAPI) (Sigma).
4. Calcofluor (Sigma).(Also known as “Flourescent Brightener 28.”)
5. Replica plating block.
6. Whatman filters, no. 1, 150 mm. (It is not necessary to sterilize filters from a
new box.)
7. Hemocytometer.
8. Light microscope equipped with a 20× long-distance working objective.
9. Fluorescence microscope.
3. Methods
3.1. EMS Mutagenesis
Optimization of the mutagenesis procedure is an empirical process. Prelimi-
nary mutagenesis studies should be performed in order to find the experimen-
tal conditions that give the highest number of potentially interesting mutants
with a reasonable level of survival (not <10%). For example, a good mutage-
nesis procedure using wild-type cells should give approximately one DNA
damage checkpoint mutant/1000 surviving cells, with a survival rate of about

10–20%. EMS has been previously used with much success for checkpoint
mutant screens in fission yeast, although alternative mutagenesis procedures
(e.g., using UV irradiation, see Note 1) should be considered, because gene
targets may differ. This may be an important consideration if the desired
mutants are rare or difficult to isolate.
1. Prepare a fresh 50-mL culture of log-phase cells (OD
600
= 0.2–0.4) growing
in YEP.
2. Collect the cells by centrifugation (~2000g) for 2 min and resuspend in 1 mL of YEP
medium containing EMS (2.5–3% v/v). Ensure the EMS is completely dissolved.
3. Incubate with shaking at room temperature for 2 h.
4. Wash the cells several times with fresh medium and plate enough cells on YEA
plates to give approx 500 colonies/plate. This should be around 5000 cells/plate,
assuming a survival rate of approx 10%.
5. Incubate the plates at 27°C. (Different permissive conditions may be required for
the isolation of thermosensitive mutants.)
3.2. Identification of S-M Checkpoint Mutants (
see
Note 2)
The HU sensitivity screen has been one of the most efficient and suc-
cessful experimental approaches for identifying new DNA structure check-
point mutants, since it provides easily definable phenotypes. HU is a
4 Martinho and Carr
powerful inhibitor of the ribonucleotide reductase enzyme that catalyzes
the rate-limiting step in the production of deoxyribonucleotides needed for
DNA replication.
1. Replica plate the mutagenized colonies from Subheading 3.1. onto two plates,
one YEA and one YEA + P with HU as follows: Press the master plate against the
replica-plating block covered with a Whatman filter. Gently remove the plate in

such way that a replica of colonies from the master plate remains on the filter.
Transfer the cells to replica plates by repeating the procedure. Remove excess
cells from the replica plates by pressing each plate against a clean filter. Incubate
at 27°C for 24–48 h.
2. Dead colonies on HU-containing YEA + P plates will appear red in color.
Phloxin B stains dead cells red, but is actively excluded from live cells. Most
of these dead colonies when observed under the microscope will contain many
highly elongated dead cells (stained red). The HU-sensitive S-M checkpoint
mutants will have a different morphology characterized by small dead cells and
no elongation.
3. From the YEA master plate, pick cells that correspond to phenotypically interest-
ing dead colonies and patch onto a new YEA plate.
4. Confirm the phenotype of these patches by replica plating again onto YEA and
YEA + P with HU. Incubate the plates at 27°C for 48 h. Discard those mutants
that do not show a reproducible phenotype (see Notes 3 and 4).
3.3. Identification of DNA Damage Checkpoint Mutants
(
see
Note 5)
The isolation of DNA damage checkpoint mutants is more difficult than the
identification of S-M checkpoint mutants, because the phenotypes observed
during the screen are not as accurate as with cells treated with HU (particularly
if UV radiation is used as the selective agent). Since most S-M checkpoint
mutants are also deficient in the DNA damage checkpoint, the following
experimental procedure should also be used to check any new S-M checkpoint
mutant previously isolated. This screen should be performed simultaneously
with the HU screen by including an extra replica plating.
1. Replica-plate the mutagenized colonies onto two plates, one YEA and one
YEA + P, as described in step 1 of Subheading 3.2. Make sure the excess of cells
is removed from both plates.

2. UV-irradiate the YEA-P plates with 200 J/m
2
.
3. Incubate at 27°C for 48 h.
4. Dead colonies on the UV-irradiated plates will appear as red “spots”. Most of
these dead colonies when observed under the microscope will contain lots of
dead cells (stained red), most of which will show some degree of elongation (see
step 2, Subheading 3.2.). In the DNA damage checkpoint mutants, this elon-
gated phenotype will be absent or greatly reduced.
Checkpoint Mutant Screen in
S. pombe
5
5. Pick from the YEA master plate those cells that correspond to phenotypically
interesting dead colonies, and patch onto a fresh YEA plate.
6. Confirm the phenotype of these patches by replica plating again onto YEA
and YEA + P media. Irradiate the YEA + P plates, and incubate at 27°C for
48 h. Discard any mutants that do not show a reproducible phenotype (see Notes
3 and 4).
3.4. Survival Analysis
In order to have a clear picture of the nature of the mutants isolated in the
screen, it is useful to determine their survival response to DNA-damaging
agents and HU, and to do a microscopic analysis of cell morphology. This
information will help to classify the mutants into groups, and identifies inter-
esting and desired phenotypes.
3.4.1. HU Survival Curves (
see
Note 6)
1. Determine the cell number of a fresh exponentially growing culture using a
hemocytometer.
2. Dilute to a cell density of ~5000 cells/mL in YEP.

3. Add HU to the diluted cell culture to a final concentration of 10 mM.
4. Incubate the culture at 30°C, take a 100-µL sample at different time-points (0, 1,
2, 3, 5, 7, 10 h) and plate onto YEA plates.
5. Incubate at 27°C for 72 h.
6. Count the colonies, and calculate the percent survival by comparing with the
time-zero control plate.
3.4.2. UV Survival Curves (
see
Notes 7 and 8)
1. Follow steps 1 and 2 of Subheading 3.4.1.
2. Plate 100-µL aliquots of the diluted cell culture onto each of 16 YEA plates
(500 cells/plate).
3. UV-irradiate the plates using the following doses: 0, 25, 50, 100, 150, 200, 250,
and 300 J/m
2
. All UV treatments should be done in duplicate.
4. Incubate the plates at 27°C for 72 h.
5. Count the number of colonies, and calculate the percent survival by comparing
with the nonirradiated control plates.
3.5. Microscopic Analysis
The morphology of the mutant cells and their nuclei after exposure to HU
can be studied using the DNA-specific fluorescent dye DAPI and an additional
dye, calcofluor, that stains material of the septum. The cells are then examined
by fluorescence microscopy to determine their morphology.
1. Take cells from an exponentially growing culture, and incubate in YEP contain-
ing 20 mM HU at 30°C.
6 Martinho and Carr
2. Take 100-µL samples of cells at 2, 6, and 18/24 h.
3. Collect the cells by centrifugation, wash once in water, resuspend in 10 µL water,
and fix in 200 µL of methanol.

4. Spot 10 µL of the fixed sample onto a glass slide, and air-dry for 5 min.
5. Pipet onto a cover slip 5 µL of a water, containing DAPI stain (0.1 µg/mL) and
calcofluor (0.5 µg/mL), and gently press against the dried fixed cells on the
glass slide.
6. Examine the cells using a fluorescent microscope, and determine the percentage
of each phenotype (cut and elongated) for each sample (see Note 9).
4. Notes
1. Alternative mutagenesis protocol using UV radiation:
a. Prepare a fresh culture of log phase cells (as described in step 1, Subheading 3.1)
b. Plate enough cells onto YEA plates to give ~500 cells per plate surviving
mutagenesis (1000-2000 cells per plate assuming a survival rate close to
25–50%).
c. Make sure the surface of the plate is well dried, remove the lid and UV
irradiate. The UV dose for wild-type cells is ~300 J/m
2
.
d. Incubate the cells as described in step 5 of Subheading 3.1.
2. Since the most obvious screens are already very close to saturation, any attempt
to isolate new genes involved in the DNA structure checkpoint response should
be designed with great care, and specific objectives and different targets decided
in order to avoid isolating previously cloned genes. For example, a cdc17 (DNA
ligase) mutant can be used in a screen comprising synthetic lethality following a
transient shift to the restrictive temperature, or a 48-h incubation at the
semipermissive temperature. The DNA ligase thermosensitive mutant when
incubated at the restrictive temperature (35.5°C) is defective in the ligation of
Okazaki fragments during replication. At the restrictive temperature, the cdc17
mutant arrests in S-phase, elongates, and slowly loses viability. This late S-phase
arrest is distinct from early S-phase arrest caused by HU. Mutations abolishing
the S-M checkpoint in a cdc17 background will make the double mutants highly
sensitive to elevated temperatures. Double mutants will rapidly become nonvi-

able after a brief incubation at the restrictive temperature (“transient temperature
sensitivity”) or a long incubation at the semipermissive temperature, since they
will enter an abortive mitotic event with unreplicated DNA, displaying a cut phe-
notype. In some aspects, screens using the cdc17 genetic background mimic the
HU mutant screen, but subtle differences exist that may be useful for the isola-
tion of new checkpoint mutants.
a. Replica plate the mutagenized cdc17 colonies onto two plates (one YEA and
one YEA + P) as described in Subheading 3.2.
b. Incubate the YEA master plate at 27°C for 48 h, and the YEA + P plate first at
35.5°C for 9 h and then at 27°C for 48 h, or incubate the YEA master plate at
27°C for 48 h and the YEA + P plate at 31.5°C (semipermissive temperature)
for 48 h.
Checkpoint Mutant Screen in
S. pombe
7
c. Identify those dead colonies on the YEA + P plate comprised of cells with a
cut phenotype. Isolate the corresponding cells from the YEA master plate,
and patch onto a new YEA plate.
d. Confirm the phenotype of these patches by replica plating again onto YEA
and YEA + P, repeating step b.
e. Discard those mutants that do not show a reproducible phenotype.
The 9-h incubation of the cdc17 mutant at 35.5°C (or 48 h at the semi-
permissive temperature of 31.5°C) reduces the viability of the single mutant,
but colonies still form. A double mutant composed of cdc17 and any S-M check-
point mutant will be nonviable and incapable of forming colonies under these
conditions. The use of DNA replication mutants like cdc20 (DNA polymerase
¡) that arrest in early S-phase (cdc17 arrests in late S-phase), is also a poten-
tially useful approach since it may uncover different aspects of the S-M check-
point pathway.
3. Genetic analysis of checkpoint mutants: To ensure that the phenotype seen in

each mutant is the outcome of a single gene mutation and not the result of the
interaction between two different genetic mutations, it is essential to backcross
each mutant three times with wild-type cells. If after this process the pheno-
type is retained, it is reasonable to assume that only one gene is responsible
for it. In addition these backcrosses have the important effect of ensuring a clean
genetic background.
4. Most mutant screens target particular genes preferentially in such a way that many
of the generated mutants may be identical (e.g., rad3 mutants constitute up to
50% of the S-M and DNA damage checkpoint mutants isolated to date). To avoid
unnecessary duplication of work by characterization of two identical checkpoint
mutants, it is recommended that mutants be crossed to one another and to known
checkpoint mutants with similar phenotypes. If the two strains used in a given
cross are allelic, then wild-type cells will not be generated from this cross. Note,
that if two different genes are closely linked, wild-type cells may be absent or
rare. However, linkage between two different nonallelic mutants with a similar
phenotype is very rare.
5. Alternative procedure: The use of a-rays in the isolation of DNA damage check-
point mutants will primarily isolate mutants deficient in G2-M arrest, since this
transition is the most critical in cells exposed to ionizing radiation. The experi-
mental procedure is essentially the same as the one described in Subheading 3.3.
for the isolation of UV-sensitive checkpoint mutants. A a-ray dose of approxi-
mately 1000–1500 Gy is required.
6. An alternative HU survival test: the spot test.
a. Determine the cell density of an exponentially growing culture using
a hemocytometer.
b. Dilute each culture to four different concentrations (10
7
, 10
6
, 10

5
, and 10
4
cells/mL) in rich medium.
c. Make three YEA + P plates containing the following concentrations of HU: 3,
5, and 7.5 mM.
8 Martinho and Carr
d. Spot 2 µL of each diluted strain onto YEA + P plates containing the different
concentrations of HU so that an increased dilution of the same strain is spot-
ted across the plate. Different cell strains should be spotted in parallel lines on
the same plate, so that comparisons of their HU sensitivity can be made.
e. Incubate the plates at 27°C for 72 h.
f. Compare the levels of growth.
The spot test and the HU survival analysis described in Subheading 3.4.1.
may give different results. This is because the spot tests measures an adaptive
response to low concentrations of HU, whereas the survival curves measure a
survival response to acute exposure to high concentrations of HU.
7. Alternative procedure: a-ray survival curves. The experimental procedure is simi-
lar to the one described in Subheading 3.4.2. for testing survival to UV radia-
tion. The plates should be irradiated at the following doses: 0, 50, 100, 200, 400,
500, 1000, and 1500 Gy. If the a-ray source has a small irradiation chamber the
cells should be diluted to the correct cell density (5000 cells/mL), irradiated and
only then plated (as described in Subheading 3.4.2.).
8. Alternative procedure: EMS survival curves. The experimental procedure is simi-
lar to the one described in Subheading 3.4.1. for HU survival curves. Incubate
the cells in medium containing 2% (v/v) EMS, and take samples as described for
determining HU survival.
9. Most DNA damage checkpoint mutants become sensitive to HU at high concen-
trations or after long incubations, but under standard treatment conditions, they
have a normal checkpoint response and are not particularly sensitive to HU. Non-

checkpoint DNA repair mutants, when incubated with DNA-damaging agents
die with a highly elongated phenotype, because they are unable to repair the DNA
damage. Some extremely sensitive DNA repair mutants die with no elongation at
normal doses of mutagens. This is because they cannot undertake transcription.
At very low concentrations of DNA-damaging agents, such mutants will display
a highly elongated phenotype.
Acknowledgment
We wish to thank Nicola Bentley for helpful comments.
References
1. Murray, A. W. (1992) Creative blocks: cell cycle checkpoints and feedback con-
trols. Nature 359, 599–604.
2. Hartwell, L. and Weinert, T. (1989). Checkpoints: controls that ensure the order
of cell cycle events. Science 246, 629–634.
3. Enoch, T., Carr, A. M., and Nurse, P. (1992) Fission yeast genes involved in cou-
pling mitosis to completion of DNA replication. Genes Dev. 6, 2035–2046.
4. Saka, Y. and Yanagida, M. (1993) Fission yeast cut5, required for S-phase onset
and M-phase restraint, is identical to the radiation-damage repair gene rad4
+
. Cell
74, 383–393.
Checkpoint Mutant Screen in
S. pombe
9
5. Kelly, T., Martin, G. S., Forsburg, S. L., Stephen, R. J., Russo, A., and Nurse, P.
(1993) The fission yeast cdc18
+
gene product couples S-phase to START and
mitosis. Cell 74, 371–382.
6. Al-Khodairy, F. and Carr, A. M. (1992) DNA repair mutants defining G
2

check-
point pathways in Schizosaccharomyces pombe. EMBO J. 11, 1343–1350.
7. Al-Khodairy, F., Fotou, E., Sheldrick, K. S., Griffiths, D. J. F., Lehmann, A. R.,
and Carr, A. M. (1994) Identification and characterisation of new elements
involved in checkpoint and feedback controls in fission yeast. Mol. Biol. Cell 5,
147–160.
8. Sachez, Y., Wong, C., Thoma, R. S., Richman, R. Wu, Z., Piwnica-Worms, H., et
al. (1997) Conservation of the Chk1 checkpoint pathway in mammals: linkage of
DNA damage to Cdk regulation through Cdc25. Science 277, 1497–1501.
DNA Repair of
C. elegans
11
2
11
From:
Methods in Molecular Biology, Vol. 113: DNA Repair Protocols: Eukaryotic Systems
Edited by: D. S. Henderson © Humana Press Inc., Totowa, NJ
Isolating Mutants
of the Nematode
Caenorhabditis elegans
That Are Hypersensitive to DNA-Damaging Agents
Phil S. Hartman and Naoaki Ishii
1. Introduction
The nematode Caenorhabditis elegans has gained widespread popularity for
use in addressing many biological problems, particularly those relating to
development (for brief topical reviews, see 1–5; for comprehensive treatises,
see 6–10). This can be attributed to both inherent properties of the organism as
well as the collegiality extant within the “worm community.” With respect to
the former, C. elegans is extremely easy to grow in the laboratory (animals are
typically propagated on agar-filled Petri dishes seeded with the bacterium

Escherichia coli) and possesses a short generation time (3 d at 20°C). The
system is genetically robust, with the availability of thousands of mutants as
well as the existence of a physical map whose sequencing (over 82 Mb fin-
ished at present) is scheduled for completion in 1999. Developmental studies
have been advantaged by the animal’s transparent nature, facilitating complete
elucidation of C. elegans’ largely invariant cell lineage.
The collegiality of the worm community is manifested as follows:
1. There is a Caenorhabditis Genetics Center (University of Minnesota; E-mail
) that maintains many stocks and freely disseminates
them on request.
2. Investigators frequently exchange information prior to publication via the infor-
mal The Worm Breeder’s Gazette (E-mail for subscrip-
tion particulars).
3. An electronic news group exists for discussion and announcements related to C.
elegans (to subscribe by E-mail, send the message “subscribe CELEGANS” to
12 Hartman and Ishii
), allowing individuals to share information readily as
well as solicit widespread input to queries.
4. Several Web sites may be accessed (e.g., elegans.swmed.edu, www.dartmouth.edu/
artsci/bio/ambros/protocols.html, probe.nalusda.gov:8000/acedocs/allace.html)
in order to obtain protocols, various literature, and sequence information.
With respect to the processing and consequences of DNA damage in C.
elegans, several areas have received particular attention (reviewed in 1). These
include the developmental regulation of DNA repair, the lethal and mutagenic
effects of cosmic radiation (as it relates to long-term human travel in space),
and the effects of DNA damage on cellular and organismal aging.
Two protocols for isolating mutants of C. elegans hypersensitive to DNA-
damaging agents are described below. The first protocol is analogous to the rep-
lica-plating methodology developed by the Lederbergs (11), although it is
considerably more labor-intensive (cf. Chapter 6). It was developed by Hartman

and Herman (12), who screened over 6400 clones to isolate nine radiation-sensi-
tive (rad) mutants. In brief, individual second-generation progeny (F
2
s) of
mutagenized animals are placed in a first set of separate wells (“rescue wells”) of
a microtiter plate and allowed to reproduce. They are then transferred to fresh
wells (“treatment wells”) and insulted with a DNA-damaging agent under condi-
tions sublethal to wild type. Several days after transfer, the second set of wells is
examined; in them, candidates will have produced either very few offspring or a
preponderance of progeny arrested at embryonic or early larval stages. Candi-
dates can then be propagated and retested using animals from the rescue wells.
The second protocol, termed “embryo rescue,” is peculiar to C. elegans and
has as a primary advantage the fact that “replica plating” is not necessary. It is
made possible because the “eggshell” of developing embryos is impervious
to most chemicals, including many DNA-damaging agents. Thus, exposure
to the toxic chemical may kill the mutant itself, but its in utero progeny will
survive, allowing propagation of putative mutants. In this procedure, the F
2
s
of mutagenized animals are incubated for several hours in a solution contain-
ing a relatively high drug concentration. They are then plated on Petri dishes en
masse. Wild-type animals survive this treatment and begin movement within
minutes after plating. Conversely, drug-sensitive mutants die and are therefore
immobilized. The latter are plated on individual drug-free Petri dishes and their
progeny retested. This protocol has been employed successfully by one of us to
isolate two methyl viologen-sensitive mutants, mev-1 and mev-2, from about
15,000 F
2
progeny of ethyl methanesulfonate- (EMS) treated animals (13).
2. Materials

The following reflect the materials specifically employed in the authors’
laboratories. More varied and extensive descriptions of the materials necessary
DNA Repair of
C. elegans
13
to propagate C. elegans are readily available (7,8; www.dartmouth.edu/artsci/
bio/ambros/protocols.html).
1. Wild-type C. elegans, strain N2 (Caenorhabditis Genetics Center, University
of Minnesota).
2. Dissecting microscope: Animals can be readily observed with transmitted light
at 20× magnification.
3. Although animals can be grown in liquid, the methods described below employ a
solid medium such as MYOB: 5.9 g of stock, 20 g of agar/L of H
2
O. Stock is: 55
g Tris-HCl, 24 g Tris-OH, 310 g Bacto-peptone, 800 mg cholesterol, 200 g NaCl.
This medium should be autoclaved, cooled to 55°C, and either poured into 35-,
60-, or 100 mm Petri dishes or pipeted into 24-well microtiter dishes (see Notes
1 and 2). After the medium has solidified, the surface is inoculated with an
overnight culture of E. coli OP50, a uracil auxotroph (available from the
Caenhorbaditis Genetics Center, University of Minnesota), and incubated at 20°C
for at least 12 h before inoculation with nematodes. A single drop is sufficient
bacterial inoculum and can be spread with a sterile glass spreader on Petri dishes.
4. M9 buffer: 5.8 g Na
2
HPO
4
, 3 g KH
2
PO

4
, 0.5 g NaCl, 1 g NH
4
Cl/L of H
2
O.
5. 32-Gauge platinum wire (ca. 1.5 cm in length) affixed to a handle (e.g., one
designed for bacterial inoculations) is used to transfer individual animals. The
wire should be flame-sterilized before a transfer is effected. With some practice,
individual or small groups of animals (visualized under the microscope) can be
scooped off the surface and transferred to another. Care should be taken not to
gouge the agar’s surface, since the animals will burrow. Typically, two to three
animals are transferred from one plate to another for stock maintenance. Most
strains of C. elegans will starve the bacterial lawn within 1 wk of transfer,
although stocks need to be transferred only once every several weeks. Animals
are typically grown at 20°C, with 15° and 25°C the permissive and restrictive
temperatures for temperature-sensitive mutants.
3. Methods
3.1. Replica Plating
Although a number of mutagens have been employed successfully with C.
elegans (reviewed in 14), EMS is most commonly used. The following is a
modification of the protocol recently reviewed by Anderson (14).
1. To obtain semisynchronous cultures of wild-type nematodes for EMS treatment,
starve the plates of bacteria for between 1 and 2 wk before usage. Such plates will
contain a few geriatric adults and many young (L1 and L2) larvae.
2. Wash these off the plate in M9 buffer, and pellet in a clinical centrifuge.
3. Using a micropipet or Pasteur pipet, inoculate the pellet of worms onto a 100-mm
Petri dish seeded with bacteria. The nematode inoculum should be small
enough (ca. 100–500) such that the bacterial lawn does not become starved.
Incubate for 2 d.

14 Hartman and Ishii
4. Wash the plate (now containing primarily fourth-stage [L4] larvae and young
adults) with 2 mL of M9 buffer, and add the contents to 2 mL of M9 buffer to
which 10 µL of EMS was previously dissolved. After 4 h at 20°C, wash the
worms once in M9, and plate on seeded 100-mm Petri dishes at a density of
~10–20/plate. Incubate at 20°C for 5–7 d.
5. Transfer individual second-generation (F
2
) L4s to individual agar-filled wells of
24-well microtiter plates, which serve as “rescue” wells. Such animals should be
abundant between d 5 and 7 after mutagenesis.
6. After 24 h, transfer the F
2
s (now egg-laying adults) to a second “treatment”
well and immediately expose to a DNA-damaging agent. Both 254 nm UV
radiation and methyl methanesulfonate (MMS) have been employed success-
fully in such mutant hunts. UV radiation can be imposed (with the lids off !) by
a single germicidal fluorescent 15-W bulb that produces approx 1 J/m
2
/s at a
distance of 55 cm. MMS should be added to the molten agar to achieve a final
concentration of 0.1 mM immediately before these wells (but not the rescue
wells) are poured.
7. After 72–96 h, examine the treatment wells. The majority will contain
nonmutants and will have >50 F
3
s and F
4
s ranging in size from embryos to
adults. Putative mutants will be signaled by wells containing either very few

animals or a preponderance of animals arrested at embryonic or early larval
stages. Most of these are not hypersensitive to the DNA-damaging agent.
Instead, they contain a mutation in some essential gene unrelated to DNA
damage tolerance. These are evident from inspection of the rescue wells, which
will contain a similar distribution of animals as in treatment wells. Only those
clones with robust growth in the rescue well, but impaired growth in the treat-
ment well are worthy of retesting.
8. Candidates should be retested as above. With EMS-mutagenized populations,
approx 1% of the clones will pass the first screening. Of these candidates, over
80% will prove to be false-positives (see Note 3).
3.2. Embryo Rescue
1. Treat a population of wild-type animals with EMS as described in steps 1–4 of
Subheading 3.1.
2. Seven days after mutagenesis, wash the F
2
animals off the Petri dishes with M9
buffer, and incubate in 30 mM methyl viologen (or some other chemical DNA-
damaging agent) for 4 h at 20°C.
3. After this treatment, wash the animals free of methyl viologen and spot in the
middle of a 100-mm Petri dish containing a lawn of E. coli.
4. Twenty-four hours after plating, most animals will have recovered and crawled
away from the center of the plate. Transfer the carcasses of dead animals at the
center of the plate onto individual 35-mm or 60-mm Petri dishes containing an E.
coli lawn on MYOB.
5. After 3 d, the in utero embryos (resistant to the chemical by virtue of their imper-
vious eggshells) should have developed into gravid adults. Retest several from
DNA Repair of
C. elegans
15
each plate by transferring them to seeded MYOB plates impregnated with 0.2

mM methyl viologen.
6. Inspect these dishes after 3–4 d. Most wild-type embryos will have developed
into L4 larvae or adults. Conversely, mev mutants will have arrested as L1 or L2
larvae (see Note 3).
4. Notes
1. Although the microtiter dishes are “disposable,” they can be reused. To do so, the
plugs of agar should first be removed with a spatula before they desiccate. The
dishes should then be washed several times in warm, soapy water, soaked over-
night in 1% sodium hypochlorite (common bleach diluted 1:5), and rinsed sev-
eral times with deionized water. The wells can then be refilled with agar. If
bacterial or fungal contamination become problematic, the microtiter dishes can
be exposed to UV light for several hours before the agar is poured. In this event,
the lids should be removed, and the light source positioned such that, as much as
possible, the sides of the wells are exposed directly to the light (UV is poorly
penetrant through plastic).
2. It is important that no condensation is present in the microtiter plates, since nema-
todes may crawl from well to well. It is for this reason that protocols employing either
96-well microtiter dishes or liquid culture have not proven successful in our hands.
3. Once mutants are isolated, they may be analyzed as explained in ref. (12). In
addition, as with other mutants in C. elegans, the genes may be cloned by trans-
formation rescue (15) once they have been mapped reasonably precisely. Owing
to the alignment of the genetic and physical maps in C. elegans, precise mapping
allows the investigator to employ YACs and cosmids corresponding to the
genetically defined region. In addition, knowledge of the DNA sequence gained
from the sequencing project can provide the investigator hints concerning spe-
cific DNA sequences that may encode the gene.
References
1. Hartman, P. S. and Nelson, G. A. (1997) Processing of DNA damage in the nema-
tode Caenorhabditis elegans, in DNA Damage and Repair: Biochemistry, Genet-
ics and Cell Biology, vol. 1 (Nickoloff, J. A. and Hoekstra, M. F., eds.), Humana,

Totowa, NJ, pp. 557–576.
2. Jacobson, M. D., Weil, M., and Raff, M. C. (1997) Programmed cell death in
animal development. Cell 88, 347–354.
3. Kornfeld, K. (1997) Vulval development in Caenorhabditis elegans. Trends
Genet. 13, 55–61.
4. Polani, P. E. (1996) Developmental asymmetries in experimental animals. Neurosci.
Biobehav. Rev. 20, 645–649.
5. Hodgkin, J., Plasterk, R. H. A., and Waterston, R. H. (1995) The nematode
Caenorhabditis elegans and its genome. Science 270, 410–414.
6. Riddle, D., Blumenthal, T., Meyer, M. J., and Priess, J. R. (eds.) (1997) C. elegans
II. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
16 Hartman and Ishii
7. Epstein, H. F. and Shakes, D. C. (eds.) (1995) Caenorhabditis elegans: Modern
Biological Analysis of an Organism. Methods in Cell Biology, vol. 48, Academic,
New York.
8. Wood, W. B. (ed.) (1988) The Nematode Caenorhabditis elegans. Cold Spring
Harbor Laboratory Press, Cold Spring Harbor, NY.
9. Zuckerman, B. M. (ed.) (1980) Nematodes as Biological Models. Behavioral and
Developmental Models, vol. 1, Academic, New York.
10. Zuckerman, B. M. (ed.) (1980) Nematodes as Biological Models. Aging and Other
Model Systems, vol. 2, Academic, New York.
11. Lederberg, J. and Lederberg, E. M. (1952) Replica plating and indirect selection
of bacterial mutants. J. Biol. 63, 399.
12. Hartman, P. S. and Herman, R. K. (1982) Radiation-sensitive mutants of
Caenorhabditis elegans. Genetics 102, 159–178.
13. Ishii, N., Takahashi, K., Tomita, S. Keino, T., Honda, S. Yoshino, K., et al. (1990)
A methyl viologen-sensitive mutant of the nematode Caenorhabditis elegans.
Mutat. Res. 237, 165–171.
14. Anderson, P. (1995) Mutagenesis, in Caenorhabditis elegans: Modern Biological
Analysis of an Organism, in Methods in Molecular Biology, vol. 48 (Epstein, H.

F. and Shakes, D. C., eds.), Academic, New York, pp. 31–58.
15. Mello, C. and Fire, A. (1995) DNA transformation, in Caenorhabditis elegans:
Modern Biological Analysis of an Organism, in Methods in Molecular Biology,
vol. 48 (Epstein, H. F. and Shakes, D. C., eds.), Academic, New York, pp. 452–482.
Drosophila
DNA Repair Mutants 17
3
17
From:
Methods in Molecular Biology, Vol. 113: DNA Repair Protocols: Eukaryotic Systems
Edited by: D. S. Henderson © Humana Press Inc., Totowa, NJ
Isolating DNA Repair Mutants
of
Drosophila melanogaster
Daryl S. Henderson
1. Introduction
The fruitfly Drosophila melanogaster offers numerous advantages as a
metazoan model for genetic dissection of conserved biological processes, such
as DNA repair. Its ease of culture, short generation time, small number of link-
age groups (2n = 8), and giant polytene chromosomes, combined with a wealth
of morphological mutants and chromosomal variants (1) accumulated over 90
years and now cataloged in FlyBase ( or http://
www.ebi.ac.uk/flybase/), make it a powerful and versatile system for genetic
analysis (2). D. melanogaster also has emerged as one of the best multicellular
eukaryotes in which to disrupt genes by transposon mutagenesis for the pur-
pose of molecular cloning (3–5), and to study cloned gene functions by trans-
formation (6). Cytological studies of flies also have reached new levels of
sophistication in keeping with recent advances in microscopy, probe technol-
ogy, and electronic imaging (7,8).
The use of Drosophila in mutagenesis research dates back more than 70

years to H. J. Muller’s momentous discovery of the mutagenic action of X-rays
(9,10). His work, together with that of L. J. Stadler on maize (11), effectively
ushered in the field of radiation genetics. In the 1940s, Auerbach and Robson
used Drosophila to demonstrate unequivocally that chemicals, too, can have
mutagenic effects (12,13). An historical account of their work and of mutation
research in general can be found in ref. (14). Beginning in the early 1970s, the
scope of mutation research in Drosophila was broadened by the isolation of
mutants potentially deficient in DNA repair. Such mutagen-sensitive (mus)
mutations render embryos and larvae hypersensitive to the lethal effects of
DNA-damaging agents. The first mus mutations were recovered on the X chro-
18 Henderson
mosome in screens that employed methyl methanesulfonate (MMS) as a selec-
tive agent (15–17). Subsequent screens using a variety of mutagens identified
mus mutations on chromosomes 2 and 3 (18–20) and brought to more than 30
the number of mus genes documented in Drosophila (1). The notion that mus
mutations disrupt DNA repair-related genes has since been confirmed by bio-
chemical assays (21) and more recently by molecular cloning (see Table 1).
Screens for mutants in Drosophila usually target a specific chromosome—
either the X, the 2
nd
, or the 3
rd
—unlike those in other organisms, which typi-
cally screen entire genomes at a time (see Chapters 1, 2, 4–6). Such specificity
is possible with flies because of the existence of balancer chromosomes. Bal-
ancer chromosomes carry multiple inversions that suppress meiotic recombi-
nation and dominant genetic markers that allow them to be traced through
successive generations. The two major autosomes, chromosomes 2 and 3, each
constitutes ~40% of the euchromatic part of the genome, whereas the X chro-
mosome makes up nearly all of the remaining ~20%. Chromosome 4 is so

small, accounting for only ~1% of the euchromatic genes, that systematic
screens for 4
th
chromosome mutants have not been considered worth the effort.
Table 1
Cloned Mutagen-Sensitive Genes
Drosophila gene
a
Homolog
b
Reference
mei-9 XPF (22)
mei-41 ATM (23,24)
mus101 rad4/cut5 R. M. Raupp, J. M. Axton,
S. pombe D. M. Glover, and D. S. Henderson
(unpublished results)
mus205 REV3 (25)
S. cerevisiae
mus209 PCNA (26)
mus308 ? (27)
(Novel protein
with putative helicase
and polymerase domains)
mus309 Ku70 (28)
a
X-linked mus mutants are numbered beginning with 101, 102, and so forth. Those on chro-
mosome 2 are numbered 201, 202, and so on for chromosome 3. Two X-linked mutagen-sensi-
tive loci, mei-9 and mei-41, are exceptions. They carry the designation mei, for meiotic, instead
of mus because the first mutant alleles of these genes were recovered in screens for meiotic
abnormalities (29), and later found to be mutagen-sensitive (16,17,30). With the exception of

mus308, mutants in all of these genes were isolated on the basis of sensitivity to MMS; mus308
mutants are preferentially sensitive to crosslinking agents (e.g., HN2).
b
Human homolog except where indicated.
Drosophila
DNA Repair Mutants 19
In general, X chromosomal screens are faster and easier than autosomal
screens, a point made clear in Subheading 3.2.
This chapter describes strategies for isolating mus mutants on the X and 2
nd
chromosomes. Chromosome 3 mus mutants can be isolated by introducing a
simple modification to the scheme used to isolate mutants on chromosome 2. In
addition, both screens are designed to permit identification of temperature-sensi-
tive (ts) mutants. Subheading 3.1. describes how to induce germline mutations
by feeding ethyl methanesulfonate (EMS) to adult males. Males thus treated are
mated to appropriately genetically marked strains to establish, after a series of
crosses, a collection of lines in which each line potentially carries a unique EMS-
induced mutation on the X or 2
nd
chromosome (Subheading 3.2.). Larvae from
each line are then tested for hypersensitivity to one or more DNA-damaging
agents. These tests are done in such a way that each mutagen-treated culture
contains potentially mutagen-sensitive larvae together with mutagen-insensitive
(mus
+
) siblings, the latter serving as an internal control. Since these two classes
of flies are readily distinguishable from one another by their phenotypic markers
(see Subheading 3.2.), absence of the first class in any mutagen-treated culture
indicates a putative mutagen-sensitive strain. Putative mutants are then retrieved
from a stock collection for retesting and further characterization.

2. Materials
2.1. Fly Strains
1. Isogenic line (see Note 1) carrying a visible marker (or markers) appropriate for
the chromosome to be screened: e.g., w (white) for the X chromosome; b pr cn
(black body, purple eyes, cinnabar eyes) for chromosome 2; st (scarlet eyes) or
red e (red Malpighian tubules, ebony body) for chromosome 3 (see Note 2).
Before undertaking a screen, the line should be tested to ensure it is not already
hypersensitive to mutagens.
2. Attached-X stock, e.g., C(1)DX,y f/Y (referred to here as X^X/Y), for screening
the X chromosome. Balancer chromosome stock, e.g., Gla/CyO for screening
chromosome 2 or Ly/TM3,Sb for screening chromosome 3. (See Note 3.)
2.2. Mutagenesis
1. 25 mM EMS (e.g., Sigma, St. Louis, MO) in 1% v/v aqueous sucrose. Prepare
fresh. EMS is listed as methanesulfonic acid ethyl ester in Sigma’s catalog.
The density of EMS is 1.17 g/mL. Store the bottle wrapped in parafilm at 4°C.
Handle with gloves in a fume hood.
2. Denaturing solution: 1 M NaOH, 0.5% v/v thioglycolic acid. Prepare fresh.
3. Glass or plastic fly culture bottles.
4. Whatman paper filters (e.g., no. 4) cut as circles to fit tightly inside the bottom of
the bottle.
5. 2-mL Syringe with long needle (e.g., 21 gage, 1
1
/
2
inches).
20 Henderson
2.3. Mutant Screen
1. Basic equipment for Drosophila culture, such as vials, bottles, anesthetizers, dis-
secting microscope, and so forth, is required (31).
2. Mutagens (see Note 4): Nitrogen mustard (HN

2
; mechlorethamine hydrochlo-
ride; Sigma). To make a 1% (w/v) stock solution, dissolve 1 g of HN
2
(i.e., the
entire contents of a bottle) into several milliliters of 0.1 N HCl, and dilute to a
final volume of 100 mL. Dispense into 1-mL aliquots, and store at –20°C. Handle
with gloves in a fume hood. Decontaminate all pipets, glassware, and so forth,
and destroy any unwanted HN2 with denaturing solution (see item 2, Subhead-
ing 2.2.). MMS (e.g., Sigma). MMS is listed as methanesulfonic acid methyl
ester in Sigma’s catalog. Store the bottle of MMS wrapped in parafilm at 4°C.
Handle with gloves in a fume hood.
3. Denaturing solution: see item 2, Subheading 2.2.
4. Multipipeter (e.g., Eppendorf Multipette
®
and 12.5 mL Combitips, Brinkmann
Instruments, Westbury, NY).
5. Large incubators set at 22°C and 29°C (if screening for ts mus mutants).
3. Methods
3.1. EMS Mutagenesis
For historical reasons and because it is effective and relatively nontoxic to
the adult fly, EMS is the most commonly used chemical for inducing germline
mutations in Drosophila. Alternative mutagenesis procedures may be worth
considering (see Notes 5 and 6).
1. Place ~100 males (e.g., w or b pr cn) into each of several bottles containing two
layers of filter paper fitted tightly at the bottom of the bottle. Leave the flies to
starve overnight.
2. The next day, prepare 0.5–1 L of EMS denaturing solution. This should be used
to decontaminate all pipet tips, glassware, and so forth, and any spills.
3. Prepare a 25-mM EMS/sucrose solution as follows. In a fume hood and wearing

gloves, add 66 µL of EMS to 25 mL of a 1% sucrose solution. The EMS will not
go into solution right away, but will sink as droplets to the bottom of the beaker.
These droplets should be dispersed by drawing them up, along with several mil-
liliters of sucrose solution, into a 2-mL syringe and expelling the solution back
into the beaker. Repeat several times until all the EMS is in solution. To ensure a
homogeneous solution, mix using a stir bar and magnetic stirrer. (See Note 7.)
4. Using the 2-mL syringe, dispense 1–1.5 mL of EMS/sucrose solution into each
bottle containing flies. Insert the needle through the bottle stopper (or cotton
plug) taking care not to let any flies escape, and dampen the filter paper with
EMS solution. Avoid soaking the filter paper, since this may cause the flies to
stick. Allow the flies to imbibe overnight in the fume hood.
5. Decontaminate the pipet tip, syringe, and any remaining EMS solution with
denaturing solution. Use 1 vol of denaturing solution/1 vol of EMS solution.
Leave in the fume hood for 1–2 d before discarding.
Drosophila
DNA Repair Mutants 21
6. The next day, transfer the flies to bottles containing food to allow them to recover.
Tap the flies to the bottom of the EMS-treatment bottle, quickly remove the stop-
per, invert the bottle over the new food-containing bottle, and tap gently to trans-
fer the flies. Leave the flies to feed overnight. Pour denaturing solution into the
EMS-treatment bottle, and leave for 1–2 d in the fume hood.
3.2. Screen for Mutagen-Sensitive Mutants
3.2.1. X-Linked Mutants (Fig. 1)
1. Cross mutagen-fed w males (from Subheading 3.1.) with X^X/Y virgin females
(see Note 8) in bottles en masse. Use ~2–3 females for every male. Transfer the
parents to fresh bottles every 1–3 d, depending on the number of eggs laid and the
level of hatching (see Note 9). Grow these cultures at 22°C (the permissive tem-
perature). At this time set up bottle stocks of X^X/Y flies so that X^X/Y virgins
will be available for collecting at the time the F
1

males eclose.
Fig. 1. Diagram of the crossing scheme used to isolate X-linked mus mutations. See
Subheading 3.2.1. for details. The females in these crosses produce two types of
gametes: X^X-bearing ova and Y-bearing ova. The males produce both X- and Y-
bearing sperm as usual. The X chromosome that is retained in males in these crosses is
denoted simply by its marker, w. Of the four types of zygotes produced by these
matings, two are viable (w/Y and X^X/Y) and two are inviable (w/X^X and YY).
22 Henderson
2. Collect w*/Y F
1
males (where * indicates an X chromosome potentially carrying
an EMS-induced mutation [see Note 10]) and cross each one separately to 3–4
X^X/Y virgin females in vials. Label each vial, e.g., with an alphanumeric code.
Grow at 22°C. This step establishes a collection of distinct lines to be tested for
mutagen-sensitivity as described in steps 3–7. (See Notes 11 and 12.)
3. Transfer the F
2
generation of each line to vials containing fresh medium (see
Note 13), and allow the females to lay eggs for 2 d at 22°C. (See Note 14.)
4. At the end of 2 d, transfer the parents to fresh vials. Keep the second set of vials
at 22°C as stocks. (See Note 15.)
5. Prepare 0.5–1 L of denaturing solution.
6. In a fume hood and wearing gloves, prepare 0.008% w/v HN2 solution (see Note
4). To treat approx 400 vials, pipet 800 µL of 1% HN2 stock solution into a 250-
mL beaker containing 100 mL of distilled water. Mix using a stir bar or by
pipeting with a 10-mL pipet. Decontaminate all pipets, glassware, and so forth,
with denaturing solution and leave them in the fume hood for 1–2 d.
7. In a fume hood, using a multipipeter, dispense 0.25 mL of HN2 solution onto the
food surface of each 2-d-old culture (consisting of mostly embryos and a few
first instar larvae). Leave the treated cultures in the fume hood for 1 d before

transferring them to 29°C (the restrictive temperature) for the remainder of
development (~7–10 d; see Note 16).
8. Determine the male:female ratio in each vial (F
3
generation). Be sure that any late
eclosing flies are counted. Vials with no or very few males (see Note 17) and
significant numbers of females contain putative mutants belonging to one of
the following classes: ts lethal mutants (the majority caused by mutations in
essential genes having nothing to do with DNA repair), ts mus mutants, non-ts mus
mutants, or false positives. Retrieve all such lines from the stock cultures for
retesting.
9. Retest the putative mutants using the protocol described in steps 5–7, except that
for each line, set up cultures at 22 and 29°C, both with and without mutagen treat-
ment. Approximately 6–10 male-female pairs/vial (depending on fecundity) should
be sufficient for these retests. Allow the females to lay eggs for 2 d, and then trans-
fer the parents to new vials to establish a second set of cultures or “replicas.” Treat
the first cultures with mutagen, and use the replicates as untreated controls. Use at
least 3 vials/line for these retests. (see Notes 18 and 19.)
10. Determine the male:female ratio in each vial. Table 2 summarizes the pheno-
typic classes that can be expected.
11. Each confirmed mus mutant should be characterized further, e.g., by mapping the
mutation, testing for allelism with other mus mutants, testing for sensitivity to other
DNA-damaging agents, generating dose–response curves, and so forth (16,17,32).
3.2.2. Autosomal Mutants (Fig. 2)
The following protocol describes the steps necessary to isolate mus mutants
on chromosome 2. A similar procedure can be followed to isolate mutants on
Drosophila
DNA Repair Mutants 23
chromosome 3, except that 3
rd

chromosome markers and balancer chromo-
somes must be used.
1. Cross mutagen-fed b pr cn males (from Subheading 3.1.) to Gla/CyO virgin
females in bottles en masse. Grow at room temperature. (See Note 20.) Transfer
the parents to fresh bottles every 1-3 d depending on the number of eggs laid and
the level of fecundity (see Note 9).
2. Collect b pr cn*/CyO and b pr cn*/Gla F
1
males, and mate each one separately to
3-4 Gla/CyO virgin females in vials, where b pr cn* represents a 2
nd
chromo-
some potentially carrying an EMS-induced mutation (see Note 21). Label each
vial, e.g., using an alphanumeric code, to keep track of each line.
3. From each vial collect b pr cn*/ CyO male and virgin female F
2
siblings and
allow them to mate in a vial containing fresh medium. (Discard all Gla-bearing
F
2
flies.) Grow at 22°C.
4. Check the F
3
progeny for the presence of b pr cn homozygotes. These are readily
distinguishable from their b pr cn/CyO siblings; the former have black bodies
and straight wings, but the latter have wild-type body color and curly (Cy) wings.
An absence of b pr cn homozygotes in any culture containing significant num-
bers of Cy F
3
flies indicates the presence of an induced recessive lethal mutation.

Such lines should be discarded (see Notes 22 and 23).
5. Transfer the remaining lines to fresh vials (see Note 13), and allow the females
to lay eggs for 2 d.
6. At the end of 2 d, transfer the parents to new vials, and keep as stocks at 22°C.
(See Note 15.)
7. Prepare 0.5–1 L of denaturing solution.
8. In a fume hood and wearing gloves, prepare 0.06% (v/v) MMS solution. To treat
approx 400 vials, pipet 60 µL of MMS into a 250-mL beaker containing 100 mL
of distilled water. As with EMS, MMS will form droplets at the bottom of the
beaker. Disperse these droplets using a needle and syringe as described for EMS
(see step 3, Subheading 3.1.).
Table 2
Classification of Mutants
a
Mutant
22°C29°C
category + Mutagen – Mutagen + Mutagen – Mutagen
mus (non-ts) L V L V
mus
ts
VVLV
mus
tsl
LVLL
ts lethal V V L L
a
L = lethal; V = viable. Lines exhibiting a mus
tsl
phenotype must be characterized
further (e.g., by mapping) to determine if the two phenotypes (mutagen sensitivity

and ts lethality) are caused by the same mutation. Mutants of this type have been iden-
tified in Drosophila, e.g., mus209
B1
(26).
24 Henderson
9. In a fume hood, using a multipipeter, dispense 0.25 mL of MMS solution onto
the food surface of each 2-d-old culture (consisting of mostly embryos and a few
first instar larvae). Leave the treated cultures in the fume hood for 1 d before
transferring them to 29°C (the restrictive temperature) for the remainder of
development (~7–10 d; see Note 16).
10. Count the numbers of b pr cn homozygotes and b pr cn/CyO heterozygotes in
each mutagen-treated vial. Lines having no or significantly reduced numbers of
homozygotes are putative mutants belonging to one of the following classes: ts
lethal mutants (the majority caused by mutations in genes unrelated to DNA
repair), ts mus mutants, non-ts mus mutants, or false positives. (See Note 24.)
Fig. 2. Diagram of the crossing scheme used to isolate mus mutations on chromo-
some 2. See Subheading 3.2.2. for details.