PREFACE
This book contains a compendium of protocols currently in use in our laboratory,
and is an update of our previous methods book. Many of these methods did not
originate in this laboratory, but were obtained from other groups and are presented
with little or no modification. We hope that by collecting them in one volume, we will
be providing a useful service to other Drosophila groups. We would be grateful for
any corrections or suggested modifications to the protocols. Please feel free to copy
and distribute this manual as you wish.
The Rubin Lab
November 1990
Berkeley, California

This book was edited by Lynn Rubin, and was composed on a MacII running
Microsoft Word 4, and an Apple Laserwriter. The typeface is Palatino. The hot fudge
sundaes were delicious.
Work in this laboratory has been generously supported by the Howard Hughes
Medical Institute and by a grant from the National Institutes of Health as well as
postdoctoral and predoctoral fellowships from the European Molecular Biology
Organization, the National Institutes of Health, the Helen Hay Whitney Foundation,
the Jane Coffin Childs Memorial Fund Foundation, the American Cancer Society(California Division), the National Science Foundation, the Miller Institute for Basic
Research, the Lucille P. Markey Charitable Trust, and the Medical Foundation of the
University of Sydney (Australia).
The cost of production of this book was defrayed by the income from a John D.
MacArthur
Professorship.

TABLE OF CONTENTS
QUARANTINE STOCKS .............................................................................................4FLY FOOD RECIPE ......................................................................................................6
POPULATION CAGES ................................................................................................7
SOME MODIFIED DROSOPHILA TRANSFORMATION VECTORS .................10
Modified Carnegie Vectors..............................................................................11
Modified Neomycin Selection Vectors ..........................................................12
Vectors Permitting Promotion Fusions with Chloramphenicol
Acetyltransferase Transcriptional Model ................................................13sevenless Expression Casettes.........................................................................14
pCAWC Vector..................................................................................................15
HSS derivatives .................................................................................................16
G418 SELECTION .........................................................................................................17
DROSOPHILA TISSUE CULTURE ............................................................................18
CYTOLOGICAL EXAMINATION OF POLYTENE CHROMOSOMES...............23
CHROMOSOME IN SITU HYBRIDIZATION USING BIOTIN LABELEDPROBES ..............................................................................................................26
IN SITU HYBRIDIZATION TO EMBRYO SECTIONS USING 35S
RIBOPROBES.....................................................................................................33
WHOLE MOUNT IN SITU HYBRIDIZATION WITH ANONRADIOACTIVE PROBE .........................................................................43
FIXATION AND PRETREATMENT FOR IMAGINAL DISC WHOLE
MOUNT NONRADIOACTIVE IN SITU HYBRIDIZATION.....................50
IMMUNOSTAINING DROSOPHILA EMBRYOS...................................................52
STAINING EYE IMAGINAL DISCS..........................................................................55
A. Labelling with antibodies ..........................................................................55
B. Cobalt Sulphide staining ............................................................................59
C. Basic Fuchsin staining ...............................................................................60
ANTIBODY AND COBALT STAINING OF PUPAL EYE DISCS .........................62
ß-GALACTOSIDASE DETECTION IN EMBRYOS .................................................65
ß-GALACTOSIDASE DETECTION IN IMAGINAL DISCS ..................................72
ADULT HEAD FROZEN TISSUE SECTIONS..........................................................74
ß-galactosidase activity staining .....................................................................74
Antibody staining .............................................................................................76

WHOLE MOUNT STAINING OF DROSOPHILA RETINA FOR
β-GALACTOSIDASE ACTIVITY....................................................................78
EMS MUTAGENESIS...................................................................................................80
LOCAL MUTAGENESIS BY IMPRECISE P-ELEMENT EXCISION.....................83
X RAY MUTAGENESIS ...............................................................................................86
MICROSCOPIC ANALYSIS OF THE ADULT RETINA.........................................87
SOMATIC MOSAIC ANALYSIS IN THE EYE........................................................90
PREPARATION OF ADULT DROSOPHILA FOR SCANNING
ELECTRON MICROSCOPY............................................................................92
PHOTOMICROGRAPHY ............................................................................................94
GENOMIC DNA PREPS ..............................................................................................97
RNA ISOLATION USING GUANIDINE HYDROCHLORIDE.............................99
PLASMID RESCUE FROM P[lacZ] LINES ...............................................................100
PREPARATION OF PROBES FOR FOOTPRINTING EXPERIMENTS................102
Purine cleavage sequence markers ..............................................................103
DNAaseI FOOTPRINTING .........................................................................................104
PRIMER LABELLING FOR PRIMER EXTENSION ASSAY ..................................106
PRIMER EXTENSION ..................................................................................................107
OLIGO-LABELLED DNA PROBES ...........................................................................108
NORTHERN BLOTS USING RIBOPROBES.............................................................110
POLYCLONAL AND MONOCLONAL ANTIBODIES..........................................113
Polyclonal Antibody .........................................................................................113
Monoclonal Antibody ......................................................................................114
ALKALI PLASMID PREPS..........................................................................................122
SMALL-SCALE PLASMID DNA PREPARATION .................................................124
MAXI PLASMID PREPS ..............................................................................................126
GROWTH OF M13 PHAGE ........................................................................................128
GEL PURIFICATION OF OLIGONUCLEOTIDES ..................................................130
PURIFICATION OF OLIGONUCLEOTIDES BY THIN LAYER
CHROMATOGRAPHY (TLC) ........................................................................132
PROCEDURE FOR SITE-DIRECTED MUTAGENESIS ..........................................133
SITE-DIRECTED MUTAGENESIS .............................................................................137
DOUBLE STRANDED SEQUENCING .....................................................................138
PREPARATION AND PURIFICATION OF E. COLI β-GALACTOSIDASEFUSION PROTEINS AND THEIR USE AS IMMUNOGENS ....................139FUSION PROTEINS FOR ANTIBODY PRODUCTION .........................................142
MAKING Trp E FUSION PROTEINS FOR ANTIBODY PRODUCTION............145
GST FUSIONS FOR ANTIBODY PRODUCTION ...................................................149
SOLID PHASE RADIOIMMUNE ASSAY FOR IgG ...............................................151
PROTEIN GELS - SDS POLYACRYLAMIDE...........................................................152
WESTERN BLOTS.........................................................................................................153
ELECTROELUTION OF PROTEINS USING ISCO APPARATUS........................155
MAKING FOOTPRINTING EXTRACTS FROM DROSOPHILA HEADS...........157
HIGH EFFICIENCY ELECTRO-TRANSFORMATION OF E.COLI......................161
BROMODEOXYURIDINE (BrdU or BUdR) INCORPORATION AND
VISUALIZATION............................................................................................163
LAMBDASORB LIQUID PHAGE PREP ...................................................................165
SMALL-SCALE PREPARATION OF BACTERIOPHAGE LAMBDA DNA........166
PREPARATION OF SINGLE STRANDED DNA (for in vitro mutagenesis).......168
BACTERIOPHAGE LAMBDA LIBRARY SCREENING: SUMMARY.................169
RAPID SMALL SCALE ISOLATION OF DROSOPHILA DNA AND RNA .......185
CAT ASSAY: DROSOPHILA TRANSFORMANTS................................................187
CHROMOSOME WALKING USING COSMIDS.....................................................188
DEEP PSEUDOPUPIL IN WHITE- AND RED-EYED FLIES .................................191
MAKING SEMINAR SLIDES......................................................................................192
APPENDIX I: REAGENTS..........................................................................................193

Quarantine Stocks

Todd Laverty

QUARANTINE STOCKSThe following is a general outline of steps which should be followed when new
stocks are brought into the lab. It is important that stocks go through the entire
quarantine procedure to prevent a mite infestation. To be safe, one should orderstocks from Bloomington or Bowling Green Stock Centers whenever possible. Stocks
should never be ordered from a lab known to have mites. If you don't know if aparticular lab has mites, ask someone before you order it.
When handling quarantine stocks, it is a good idea to treat them as if there weremites present. Therefore, all stocks should be kept in vials with tight cotton plugs, notbottles. New vials should be set up shortly after the adults eclose, and mostimportantly, quarantine stocks must be kept isolated from all other stocks.
1) New stocks must be brought directly to an isolated spot. Do not bring them near
any lab stock.
2) At a quarantine microscope, transfer the adults to a fresh vial (P0) and keep theoriginal vial for a few days.
3) When it is obvious that the P0 vial is going fine, discard the adults and the originalvial.
4) Shortly after the adults eclose from the P0 vial, transfer them to a fresh vial (F1).
5) Keep the stock isolated for two more generations (F1 and F2).
6) After you have set up the F2 vial, check the P0 vial for mites.
7) When the adults eclose from the F2 vial, put them in a fresh vial and check the P0
and F1 vial for mites.
8) If no mites are found, the stock may be taken out of quarantine.
9) If mites are found at any time or suspected of being in a stock, transfer the adults
from the suspect vial into a vial containing a Tedion strip (strips are made bysoaking filter paper in a 0.5% (w/v) solution of Tedion in acetone and then allowed
to air dry).
10) Autoclave the old vial.
11) Every two days, transfer the adults to a new vial containing Tedion and autoclave
the old vial. Continue this for one week.
12) Place adults in a new vial containing Tedion and follow steps 3-7 as outlined above.
The only exception is that you allow two additional generations (F3 and F4) inisolation.

6

Quarantine Stocks

The Curator of Drosophila Stocks
Dept. of Biology
Jordan Hall A503
Indiana University
Bloomington, IN 47405(812) 855-5783(812) 855-2577 (FAX)
Kathy Matthew, Curator (812) 855-5782
Mid-America Drosophila Stock Center
Dept. of Biological Sciences
Bowling Green State University
Bowling Green, OH 43403(419) 372-2631

7

Todd Laverty

FLY FOOD RECIPE
1) Prepare the following mixture:watermolasses, unsulfured
agar
cornmeal
yeast, baker's

14,000 ml
1,000 ml
148 g
1,000 ml412 g

Note: If the cornmeal and yeast are mixed together dry, you will havefewer problems dissolving the yeast in the water.
2) Boil for 10 minutes.
3) Turn off the heat and slowly add:
tegosept(10% (w/v) methyl p-hydrobenzoate
in 95% ethanol)
propionic acid

225 ml80 ml4) Mix well and begin pumping.
5) This recipe yields approximately 10,000 ml of food, enough for 1,000 vials or 200
bottles.
References: Wirtz, R. A. and H. G. Semey, DIS 58: 176-180.

8

Todd Laverty
POPULATION CAGES
Starting a population cage:
A new cage should be started from 80 synchronous bottles. In a few days the flies
should become acclimated to the cage and start laying a substantial number of eggs.
Every day the flies in the cage should be fed two to four collecting plates covered
with a thin layer of yeast paste. An empty weigh boat with a couple of moist paper
towels in it will help keep the cage humid, and should also be changed daily.
Setting up fly boxes:
Fly boxes are made from plastic food storage boxes (9" x 6") with a nitex covered
opening (4" x 2") cut in the top. A monolayer of absorbent cotton is placed on thebottom and the following solution is poured evenly over it:
1 box
water

200 mlpropionic acid

0.13 ml

phosphoric acid

0.75 mllive yeast

54g

sucrose27g

Eggs for the fly box are collected in the following manner:
1) Collect eggs on freshly yeasted plates overnight.
2) Under gently running water brush the eggs off the plate into a nitex sieve.
3) Wash the eggs thoroughly with water to remove any excess yeast.
4) Fold a filter paper circle (9 cm) into a cone and place it in a funnel. Using 70%
ethanol, rinse the eggs off the nitex into the filter paper cone.
5) Place a filter paper circle (9 cm) on some paper towels. Transfer some eggs onto thecircle. Then using a small brush and 70% ethanol spread the eggs uniformly over thecircle in a monolayer.
6) Repeat step 5 until all the eggs collected are on the filter paper circles.

9Population Cages

7) Allow the filter paper circles to briefly air dry then place on the moist cotton, twocircles per box. Cover the box securely.
8) Incubate the boxes at 25°C. Flies should eclose in about 10 days.
Once the flies eclose they will only live for a day or two in the fly box. The box can beeasily emptied in the following manner:
1) Hold the box upside down and place the CO2 outlet next to, but not through, thenitex until all the flies stop moving.
2) With the box still upside down, remove the cover and dump the flies onto an open
manila folder.
3) Quickly dump the flies off the folder into an empty fly bottle sitting on ice.
Maintaining the population cage:
To maintain the population cage, three fly boxes are set up to start a new cage. If
possible the boxes should be set up from one overnight collection. If this is notpossible, some effort should be made to synchronize all three boxes so that they all
eclose within 24 hours of each other.
The following is a typical schedule for a fly cage:1) On Monday start the cage with fresh flies. Between 50-100 ml of flies is desirable fornormal uses.
2) On Friday set up three fly boxes.
3) On the following Friday the cage is cleaned of old flies, washed and dried.
4) On Monday add the new flies from the fly boxes to the cleaned cage and begin thecycle again.
This method is almost impervious to mite problems since fresh flies are used every two
weeks. It is important to thoroughly clean the cage and fly boxes after each use with
hot soapy water to prevent any contaminates from being passed on.

10Population Cages

Collecting eggs for DNA or RNA extracts:
1) Eggs are collected and washed into a nitex sieve as described above.
2) Place the sieve on a flat surface and fill with 50% bleach. Let stand for 2 minutes todechorionate the eggs.
3) Rinse the eggs with 2 liters of 0.7% (w/v) NaCl in water.
4) Place the eggs in an eppendorf tube and freeze in liquid nitrogen.
5) Store at -80°C.
Egg collecting plates:
1) Prepare the following mixture:
water

2500 mlmolasses

360 mlagar

88g2) Autoclave for one hour.
3) Slowly add:
tegosept(10% (w/v) methyl p-hydrobenzoate
in 95% ethanol)

0.5 mlethyl acetate

20 ml4) Mix well and pour into plastic weigh boats (5" x 5").
5) Let cool and store at 4°C.
6) This recipe yields approximately 18 plates.

11

SOME MODIFIED DROSOPHILA TRANSFORMATION VECTORS
Modified Carnegie Vectors

D. Mismer

Modified Neomycin Selection Vector

D. Mismer

Vectors Permitting Promotion Fusions with
Chloramphenicol Acetyltransferase
Transcriptional Module
sevenless Expression Casettes

D. Mismer
D. Bowtell

pCAWC Vector

D. Bowtell

HSS Derivatives

Kevin Jones

12

Transformation Vectors

13

Transformation Vectors

14

Transformation Vectors

15

Transformation Vectors

16

Transformation Vectors

17Transformation Vectors

18

Transformation Vectors

Kevin Moses

G418 SELECTION
G418 selection
I have found that a concentration of 1 mg/ml G418 cleanly selects between one
dose of an insertion of pHSNEO and wild-type. However there is some batch variationin the effective percentage of the drug. Therefore I recommend a pilot run with a known
transformed stock and wild-type, at perhaps 0, 0.5, 1.0, 1.5 and 2.0 mg/ml G418. I also
recommend running such controls in each experimental run.
1) G418 is made up as 100 mg/ml stock solution and stored at -20°C in 5 ml aliquots.
2) Yeast-glucose Drosophila medium is made by boiling the following for 5 minutes:
10% weight/volume dry yeast
10% weight/volume glucose (sucrose works also)
1.6% weight/volume agar
3 % volume/volume nipagen (≈tegosept, 10% w/v in ethanol)
3) Bring the food to ~ 50°C in a water bath.
4) Add the G418 with a dash of bromophenol blue (in water) and mix with a flea (theblue dye both demonstrates that the mixing is complete and marks the food as it will
be poisonous to most flies).
5) Pour tubes or bottles in the normal way and store the food at 4°C.
6) Before use bring to room temperature and add a minimum quantity of wet yeast (sothat the adults will completely consume it in two days.
7) Allow the adults two days on the food and then sub on.
8) If using heat-shock inducible vectors, raise the cultures once to 37°C for 30 minutes on
the next day. This should suffice.

19

Don RioDROSOPHILA TISSUE CULTURE
Maintenance
We mainly use the Schneider line 2 cell line and the Kc cell line. Both are diploidand originally derived from embryos. Schneider L-2 grow in M3 medium (see recipe)
supplemented with 10% heat inactivated (56° C for 30 min) fetal calf serum andantibiotics (penicillin and streptomycin). Kc cells grow in D-22 medium with
antibiotics (penicillin and streptomycin) but don't require serum. Both lines grow at
25°C without CO2 and are maintained in 25 cm2 T-flasks (Corning). Routinely cells
are grown at 1-5x106 cells/ml and are split into fresh medium at a 1 to 4 or 5 dilution
every 3 days. The cells don't attach well to a solid substrate and so are resuspended by
pipeting for subculturing. No trypsin is required.
Freezing and thawing
1)

Grow cells to 3-5x106 cells/ml (log phase) in a 30-50 ml of medium in a 150 cm2
T-flask.

2)

Resuspend cells by pipeting (sterile) and transfer to 50 ml sterile polypropylene
tube. Spin in IEC table top centrifuge (at setting 5) for 1-2 min.

3)

Remove medium by aspiration and resuspend in 1.5 ml freezing medium. Use
either M3 (for L-2) or D-22 (for Kc) and 10% DMSO (Schwartz-Mann,
spectropure) and 20% H.I. FCS.

4)

Aliquot 0.5 ml cells into 2 ml nunc vials or provials. Label and transfer tofreezer box with foam inside (to allow slow cooling).

5)

Transfer to -70°C freezer overnight (or longer).

6)

Transfer to liquid nitrogen tank for permanent storage.

To thaw:
1)

Remove vial from liquid nitrogen and warm in water bath at 25°C (or room
temperature).

2)

As soon as the contents are thawed, transfer to 25 cm2 T-flask with 5-10 ml
medium.

3)

Allow cells to attach (3 or more hrs) and replace medium with a fresh aliquot.

4)

Incubate at 25°C for 3-5 days.

5)

The cells may take a while to fully recover.

20

Drosophila Tisue Culture

DROSOPHILA TISSUE CULTURE CELL TRANSFECTION
A. Solutions
1. 0.25 M CaCl2 :for 100 ml:
3.68g CaCl2 2 H2O
ddH2O to 100 ml

Sterile filter, aliquot into 15 ml polypropylene tubes. Store at -20°C.
2. 2XHEBS
NaCl
KCl
Na2HPO4
dextrose
Hepes (free acid)for 1 L:
16g
0.7g
0.4g
2g
10g

pH to 7.1 with NaOH
ddH2O to 1 L
Sterile filter, aliquot in 50 ml tubes
Store at -20°C.
NOTE: When thawing for a transfection, re-pH to 7.1 and re-sterile filter before
use.
B. Procedure
1)

Seed 5 ml medium in 60 mm dish (Falcon) with 0.2 - 0.3 ml of cell culture (58x106 cells/ml; e.g. 2-3 day old culture).

2)
3)

Incubate at 25°C at least 6 h or overnight before transfection.
Mix 10 µg plasmid DNA with 0.4 ml 0.25 M CaCl2 and add to 0.4 ml 2X HEBES
pH 7.1 dropwise with swirling in 17X100 mm polycarbonate tubes (Falcon).

4)

Incubate at room temperature for 20-30', the solution should become lightly
cloudy.

5)

Add 0.8 ml ppt/60 mm dish, swirl and incubate at 25°C.

6)

For selection: (e.g. for transformed cell lines) after 24 h: split cells 1 Æ 4 into
new medium. After another 24 h add selective drug. G-418: (for pcopneo andderivatives) at 1 mg/ml; Hygromycin (pcophyg and derivatives) at 200 µg/ml.

7)

Split cells every 7-10 days into selective medium.

8)

Grow cell lines as mixed cultures in selective medium.

21

Drosophila Tisue Culture

ECHALIER'S D-22 MEDIUM FOR Kc CELLS (HIGH PO4) - 10L - 2X.D22
1)

Dissolve in less than 500 ml
L-glutamic acid
79.4g
glycine
40.4g
Adjust to pH 7.0 with KOH pellets (about 37.5 gm; not much more, if any!)

2)

Dissolve in less than 500 ml
L-glutamic acidglycineAdjust to pH 7.0 with NaOH pellets.

138.18g70.32g

Dissolve in 250 ml or less
MgCl2 6 H2O
MgSO4 (anhydrous)

20g
36g

4)

Dissolve in ~150 ml
Na H2 PO4 H2O

8.32g

5)

Dissolve in less than 500 ml
L-malic acidsuccinic acidsodium acetate 3 H2Oglucosephenol red (sodium salt)

13.4g
1.2g
0.5g
40g
0.2g

6)

Dissolve in 1 L
CaCl2

17.8g

7)

Vitamin de Grace - 500X

40 ml

8)

Dissolve in 1 L
yeast extract

30g

9)

Dissolve in 4 L
Heat to dissolve. Cool and add slowly to media.
Lactalbumin hydrolysate
300g

3)

Mix in this order:1)

Add CaCl2 (6) to MgCl2 and MgSO4 (3) slowly with stirring on stirrer.

2)

Add K-glutamic/glycine (1)Na-glutamic/glycine (2) and Na H2 PO4 H2O (4)malic acid/succinic acid/glucose/NaOAC (5)yeast extract (8) and lactalbumin hydrolysate (9)500X graces vitamins (7) then add PEN/STREP
for 10 L 2 X - add 1.0g streptomycin sulfate
0.63 g penicillin G 1585u/mg

22

Drosophila Tisue Culture

3)
4)
5)
6)

Adjust to pH 6.7 with KOH
Adjust volume to 10 L
Filter sterilize and dispense into sterile 1L bottles.
Store at 4°C.
500X GRACE'S VITAMINS FOR ECHALIER'S D22 MEDIUM
Vitaminmg/Lthiamine hydrochloride riboflavin10

calcium pantothenate

10

pyridoxine hydrochloride

10

p-aminobenzoic acid10

folic acid10

niacin10

isoinositol (myo-inositol)

10

biotin5cholin chloride

100

23

Drosophila Tisue Culture

M3 MEDIUM - FOR SCHNEIDER LINE-2 CELLS: 10L (1X)gga-alanine15oxaloacetic acid

2.5b-alanine

2.5

phenylalanine

2.5

arginine-HCl

6..04

proline4

asparagine

3

serine3.5

aspartic acid3

threonine5

cysteine-HCl2tyrosine

2.5

glutamine6

tryptophan

1

K glutamate

71.5

valine4

Na glutamate

65.3

glucose100

glycine5bactopeptone

25

histidineisoleucine

5.5
2.5

yeastolate
CaCl2

20
7.6

leucine

4

MgSO4

21.5

lysine-HCl

8.5

Na H2PO4 H2O

7.8

methionine

2.5

BIS-Tris

10.5

Dissolve in 9 L dd H2O
Add 1 mg choline and 10 mg KHCO3
Adjust pH to 6.8 with NaOH, adjust volume to 10 L
Filter and store at 4°C
For complete medium add heat inactivated fetal calf serum and pen/strep.
To make radiolabeling medium (for 35S-met): omit methionine, yeastolate andbactopeptone.

24Todd Laverty

CYTOLOGICAL EXAMINATION OF POLYTENE CHROMOSOMESThis method of preparing polytene chromosomes is slightly different from traditional
methods. In the past chromosomes were spread by applying pressure to the slidewith one's thumb or with a vise. Described below is a modified technique of Johng K.
Lim's of the University of Wisconsin-Eau Claire, in which the chromosomes are gently
spread by a wave of stain flowing under the coverslip.
1) For the best results larvae should be grown in well yeasted media at 18°C and careshould be taken not to overcrowd the bottles.
2) On a clean siliconized slide place two drops of 45% acetic acid and a small drop (2-3mm) of lacto-aceto orcein stain.

3) In the first drop of 45% acetic acid dissect out the salivary glands from a third instar
larva.
4) Transfer the glands to the second drop of 45% acetic acid and cut off the anterior
ends of the glands. Quite often this will allow the glands to separate from themembrane and the attached fat body. If this does not occur, carefully dissect away
the fat body.
5) Transfer the "fat free" glands into the staining solution and incubate for 2-3 minutes.
6) Place a clean siliconized coverslip (18 mm square) over the glands in the stain.
7) Place the slide on several sheets of paper towel. With another sheet of paper towel
hold the edge of the coverslip to prevent it from moving.

25

Cytological Examination of Polytene Chromosomes8) Using a stiff, pointed probe, gently press once over the glands. Then gently tap thecoverslip, starting over the glands, working your way toward the edge in a spiralpattern.

9) Turn the slide over so that the coverslip is between the paper towel and the slide.
Press gently on one edge of the slide so that the stain flows to the opposite edge.

10) Turn the slide back over and again hold the edge of the coverslip with a paper
towel.
11) Using the probe, gently streak the coverslip with a rapid back and forth motion at
one edge and then gradually move to the opposite edge.

12) Blot off excess stain and examine under the microscope using phase contrast. If
spreading is not sufficient, gently restreak the coverslip.

26

Cytological Examination of Polytene Chromosomes13) To make the slide permanent, keep the slide overnight at room temperature.
14) Freeze the slide in an ethanol-dry ice bath for 60 minutes)
15) Pop off the coverslip and immediately place the slide in 95% ethanol (roomtemperature) for 2-3 minutes.
16) Place the slide in 100% ethanol (room temperature) for 10 minutes.
17) Place a drop of mounting media over the spread and cover with a 22 mm square
coverslip.
Lacto-aceto orcein stain:
Incubate a 2% (w/v) solution of "Gurr's orcein natural" in concentrated lactic acid atroom temperature for several hours, then filter. Heat, but do not boil, a 2% (w/v)
solution of "Gurr's orcein natural" in concentrated glacial acetic acid for several hours,
then filter. Combine the filtered solutions with water in a 1:1:1 ratio.
Gurr's Orcein Natural
Cat. no. 22100
Bio/Medical Specialties
Box 1687
Santa Monica, CA 90406(213) 829-2373

27

Todd Laverty

CHROMOSOME IN SITU HYBRIDIZATION USING BIOTIN LABELEDPROBES
Preparation of Chromosome Squashes:
This method of preparing polytene chromosomes for use with in situ
hybridization is very similar to the protocol described in the previous chapter. It is aneasily learned technique and with a little practice will yield consistent results.
1) For the best results larvae should be grown in well yeasted media at 18°C and careshould be taken not to overcrowd the bottles.
2) On a clean siliconized slide place two drops of 45% acetic acid and a clean
siliconized 18 mm square coverslip. On the coverslip place a small drop (2-3 mm) of
fixative( 1/2/3 mixture of lactic acid/water/glacial acetic acid).

.
3) In the first drop of 45% acetic acid dissect out the salivary glands from a third instar
larva.
4) Transfer the glands to the second drop of 45% acetic acid and cut off the anterior
ends of the glands. Quite often this will allow the glands to separate from themembrane and the attached fat body. If this does not occur, carefully dissect away
the fat body.
5) Transfer the "fat free" glands into the fixative and incubate for 4-5 minutes.
6) Pick up the coverslip and glands with a clean slide by carefully touching the slide to
the drop of fixative.
7) Place the slide, coverslip up, on several sheets of paper towel. With another sheet ofpaper towel hold the edge of the coverslip to prevent it from moving.

28

Chromosome in situ hybridization using biotin-labelled probes8) Using a stiff, pointed probe, gently press once over the glands. Then gently tap thecoverslip, starting over the glands, working your way toward the edge in a spiralpattern.

9) Turn the slide over so that the coverslip is between the paper towel and the slide.
Press gently on one edge of the slide so that the fixative flows to the opposite edge.

10) Turn the slide back over and again hold the edge of the coverslip with a paper
towel.
11) Using the probe, gently streak the coverslip with a rapid back and forth motion at
one edge and then gradually move to the opposite edge.

12) Blot off excess fixative and examine under the microscope using phase contrast. If
spreading is not sufficient, gently restreak the coverslip. On a good slide thechromosomes should appear low contrast, cytoplasmic debris should be vague,and no fixative should be flowing between the slide and the coverslip.

29

Chromosome in situ hybridization using biotin-labelled probes13) Incubate the slide over night at a temperature between 4° and 18°C. Leaving theslides at room temperature for a couple of hours before incubating them will help
when incubating at or near 4°C.
14) Place the slides in a dry ice/ethanol bath for 60 minutes. During or before this timecool down some 100% ethanol in a staining dish to -60 to -80°C.
15) Remove the slides individually from the dry ice/ethanol bath and quickly flip off
the coverslip with a razor blade and place the slide immediately into the chilledethanol.
16) Allow the ethanol to warm up to room temperature, then air dry the slides.
Chromosome squashes can be stored at this stage for several months before
hybridization.
Pretreatment of Chromosomes for Hybridization:
The chromosomes used for hybridization must be non-refractile after ethanol
dehydration (A [16]). The chromosomes on the slide should look the same at this point
as they did when the slide was made or it is not worth using. In general one slide isenough for a single probe if the chromosomes have good morphology. After the
pretreatment the slides should be hybridized within 24 hours.
1) Place the slides in 2X SSC at 65°C for 30 minutes.
2) Wash for 2 minutes in 2X SSC at room temperature.
3) Acetylate the slides as follows: Prepare 500 ml of 0.1 M triethanolamine-HCl (1Mstock pH 8.0) in a staining dish containing a magnetic stirring bar. Rapidly agitate
the solution and add 0.625 ml of acetic anhydride. Turn the stirrer off and
immediately place the slides into the solution. Incubate for 10 minutes. (Somepeople claim this step is unnecessary, but it has not been tested in this lab.)
4) Wash the slides in 2X SSC two times for 5 minutes each.
5) Ethanol dehydrate two times 5 minutes in 70% ethanol and 5 minutes in 95%
ethanol. Air dry.
6) Denature the chromosomes by incubation in freshly prepared 0.07 N NaOH for 3
minutes.
7) Repeat steps 4 and 5.

30

Chromosome in situ hybridization using biotin-labelled probes

Biotinylation of DNA:The following protocol is for 0.5 µg of DNA or less. The quality of the DNA is notimportant. Quick prep DNA and DNA fragments isolated by gel purification work
equally well as CsCl purified DNA. Probes can be stored at -20°C for several months.
1) Prepare a 1:400 dilution of a 1 mg/ml stock of DNAse I (Worthington, DPFF grade)in 10 mM Tris, pH 7.5/10 mM MgCl2 (TM10). The DNAse I stock should be
prepared in TM10 and should be stored in 20µl aliquots at -20°C.
2) Mix:

10X nick-translation bufferdATP, dGTP, dCTP mix (0.3 mM each)Bio-16-dUTP
32P-dATP (1mCi/ml)
DNA
DNAse I (1:400)
distilled water to

2.5 µl
2.5 µl
2.0 µl
1.0 µl
~ 0.5 µg
1.5 µl25 µl

add 10 U of DNA polymerase I and incubate at room temperature for 60
minutes.
3) Stop the reaction by adding 25 µl of 50 mM EDTA, pH 8.0.
4) Remove unincorporated nucleotides by separation over a P 10 spin column.
5) Check the percentage of incorporation by comparing the signals of the column andthe flow through on a Geiger counter. 5 - 10% incorporation is sufficient. Probes thatincorporated more than 40 - 50% might not do so well in the hybridization, since
oversubstitution of the Bio-16-dUTP renders the forming hybrids less stable.
6) Add 20 µg of carrier DNA (Herring Sperm or equivalent) and 5 µl of 3 M sodium
acetate. Precipitate the DNA by adding 2.5 volumes of ethanol and incubate in dry
ice/ethanol for 20 minutes or at -20°C overnight.
7) Recover the DNA by centrifugation and dry under vacuum for 2 minutes.
8) Resuspend the pellet in 75 µl of hybridization buffer.

31

Chromosome in situ hybridization using biotin-labelled probes

Hybridization and Washes:1) Denature the hybridization probe by boiling for 3 minutes and chill on ice.
2) Apply 5 µl of hybridization solution on the slides and distribute the solution over thechromosomes with a 18 x 18 mm coverslip. Try to avoid trapping air bubbles.
3) To prevent evaporation of the probe seal the edges of the coverslip with rubber
cement.
4) Incubate in a moist chamber at 58°C for 12-18 hours.
5) Place the slide in a beaker containing 2X SSC and pry off the coverslip and rubber
cement with a scalpel blade.
6) Wash the slides 3 times 15 minutes in 2X SSC at 53°C. Proceed directly with thesignal detection. It is important not to let the slide dry out during the signal
detection procedure.
Signal Detection:
The age or storage conditions of the Detek I-hrp kit can have a dramatic effect on the
kit's proficiency to detect signals. Kits that are over six months old or that have notbeen kept cold (2-8°C) can give weak or inconsistent signals. On the other hand, underideal conditions probes can be diluted 10-fold and still yield strong signals.
1) Wash the slides in 1X PBS 2 times for 5 minutes each at room temperature.
2) Wash for 2 minutes in 1X PBS/0.1 % Triton-X-100.
3) Rinse in 1X PBS.
4) Place the slides horizontally on two 5ml pipettes in a tray lined with moist paper
towels. Apply 100µl of a 1:250 dilution of the Streptavidin-biotinylated peroxidase
complex in 1X dilution buffer (all the components are supplied in the Detek I-hrp kit
by ENZO Diagnostics, Inc.) Distribute the solution by placing a 22 x 40 mm
coverslip on the slide.
5) Cover the tray with plastic wrap and incubate at 37°C for 30 minutes.
6) Float the coverslip off in a beaker of 1X PBS and repeat steps 1 through 3.

32

Chromosome in situ hybridization using biotin-labelled probes

7) Prepare a solution containing 0.5 mg/ml of diaminobenzidine in 1X PBS. Add 1/100
volume of 1% hydrogen peroxide (prepared from a 30 % stock in distilled water).
The staining solution should always be prepared fresh. Diaminobenzidine is a
carcinogen. Wear gloves when handling the solution. Diaminobenzidine can beinactivated by bleach which should be used liberally when disposing of unuseddiaminobenzidine and contaminated materials.
8) Place the slides horizontally in a tray and add 50 µl of staining solution onto eachslide, cover with a 22 x 40 mm coverslip and incubate at room temperature for 1 - 10
minutes. The staining reaction can be observed by blotting off excess solution andexamining the slide under the microscope using a low magnification phase contrast
lens (e.g. 16x).
9) Stop the reaction by dipping the slides in distilled water three times and then placethem in distilled water until staining.
10) Prepare a 4% dilution of Giemsa stain in 10 mM sodium phosphate buffer, pH 6.8.
Stain the slides for 30 seconds, rinse in water and allow the slides to air dry. It is
important not to overstain the chromosomes for this may obscure thediaminobenzidine precipitate.
11) Place a drop of mounting media over the chromosomes and cover with a 22 mm
square coverslip.
12) Examine the chromosomes using phase-contrast optics. Hybridization signals
appear as black bands. Weaker signals are brown.
Solutions:
1) 10X PBS
1.3 M NaCl
0.07 M Na2HPO4
0.03 M NaH2PO42) 10X Nick-Translation Buffer
0.5 M Tris-HCl, pH 7.5
0.1 M MgSO4
1 mM dithiothreitol (DTT)
500 µg/ml bovine serum albumin (BSA Pentax Fraction V)
3) Hybridization Buffer
0.6 M NaCl
50 mM Sodium phosphate buffer, pH 6.8
1X Denhardt's (0.02% BSA / 0.02% Ficoll / 0.02% polyvinylpyrrolidone)
5 mM MgCl2

33

Chromosome in situ hybridization using biotin-labelled probes

Reagents and Suppliers:
1) Bio-16-dUTP
Cat. No. 42811
ENZO Diagnostics, Inc.
325 Hudson St.
New York, NY 10013
1-800-221-7705
2) Detection Kit
Detek I-hrp
Cat. No. 43820
contains: Streptavidin-biotinylated peroxidase complex / 10X dilution buffer
ENZO Diagnostics, Inc.
325 Hudson St.
New York, NY 10013
1-800-221-7705
3) Diaminobenzidine
3',3' diaminobenzidine tetrahydrochloride
Cat. No. D 9015
Sigma Chemical Company
P.O. Box 14805
St. Louis, MO 63187
1-800-352-3010
4) DNAse I (Grade DPFF)
Cat. No. LS 6330
Worthington Biochemicals
Halls Mill Road
Freehold, NJ 07728
1-800-445-9603

34

Matthew Freeman

IN SITU HYBRIDIZATION TO EMBRYO SECTIONS USING 35S
RIBOPROBES
Introduction
The determination of the tissue distribution of a transcript of interest has proved to
be of great use. Originally the probes were made with tritium. Although this gave very
good localisation, the exposure times were rather long (a month was not uncommon).
Several protocols were then published using 35S as the labelling isotope, and theexposure times necessary have been cut 5 or 10-fold.
Recently, it has become possible to do non-radioactive in situs (described elsewherein this manual). These have several major advantages, the main one being the ability tolocalise the expression with single-cell resolution. However, there are cases whereradioactive in situs might still be the method of choice: they may be a bit more sensitive,
and tissues deep within the embryo seem a bit less accessible to digoxygenin-labelledprobes than to 35S labelled probes. I would try a non-radioactive whole-mount in situ
first, and if that was unsuccessful, resort to the old-fashioned way. In time, as the nonradioactive protocol is refined, this one may become obsolescent.
This protocol is very largely that of Phil Ingham.
Fixing the embryos:
1) Use as many embryos as it is practical to collect. Less than about 500-1000 is probably
much harder to work with than a gram or two.
2) Wash embryos from collection plate into a sieve with a fine enough mesh to retain theembryos. Rinse them with water to remove excess yeast.
3) Dechorionate in 50% household bleach for about 2 minutes.
4) Rinse the dechorionated embryos thoroughly with water containing 0.1% Triton X100.
5) Transfer the embryos to a tube with a tightly fitting lid that contains a 1:1 mixture of
4% paraformaldehyde in PBS1, and heptane. For about a gram of embryos I usually
use a 50ml tube, with 20ml each of formaldehyde and heptane. For fewer embryos,
scale down the sizes appropriately.
6) Shake the tube vigorously for 15 minutes. Then remove as much as possible of theaqueous (bottom) layer with a pipette, being careful not to remove the bulk of theembryos that will be at the interface.
7) Add a volume of methanol equal to the heptane layer. Shake the tube very hard forabout 30 seconds, and then allow the two phases to separate. The devitellinised
embryos will sink to the bottom of the tube, and the rest will remain at the interface
1This must be made fresh. Heat it until near boiling, and then add a drop or two of 1M NaOH. It

should go into solution within a few minutes.

35

In situ hybridization to embryo sections using 35S riboprobeswith the empty vitelline membranes. The yield at this stage is variable - anythingover 50-60% is fine; if there are fewer than that, try shaking harder and longer.
8) Remove as much liquid as possible and all the interface, leaving the embryos at thebottom of the tube.
9) The rest of the treatments are carried out in the same tube as the embryos are now in;each liquid is added to the tube, the embryos allowed to settle, and the liquid
removed with an aspirator.
10) Rinse the embryos in methanol for 2 minutes.
11) Rehydrate them as follows:
2 minutes in 9:1 methanol:4%paraformaldehyde
2 minutes in 7:3""
2 minutes in 5:5""
2 minutes in 3:7""
12) Post-fix the embryos in 4% paraformaldehyde for 20 minutes.
13) Wash once in PBS.
14) Dehydrate in an ethanol series for 5 minutes each: 30%, 50%, 70%.
15) The embryos can be stored in 70% ethanol at -20° for several weeks.
Embedding the embryos:
1) Fully dehydrate the embryos as follows:
90% ethanol, 5 minutes, twice
95% ethanol, 5 minutes, twice
100% ethanol, 3 minutes, four times2) Transfer the embryos to a glass test tube, and incubate them in xylenes, twice for ten
minutes each.
3) Incubate in a 1:1 mixture of Paraplast Plus (Polysciences):Xylenes2 at 61° for ten
minutes, twice. Keep the tubes in a hot block for the rest of the procedure.
4) Incubate in Paraplast Plus at 61° for ten minutes, twice.
5) Set up the molds3 on a hot plate set just hot enough to ensure that the wax does not
set when a little is put in the bottom of the mold.
6) Quickly dispense some of the embryos in wax into the bottom of the mold. Use apasteur pipette with the end broken off for this operation. I warm it briefly in a
2Paraplast Plus melts at 56°, and must not get hotter than 62°. Melt some in a conical flask in awater bath, and add an equal volume of pre-warmed xylenes. The solid wax takes quite a longtime to melt, and should be set up at least an hour in advance. Also set up the water bath and the
hot-block for the test tubes of embryos far enough in advance that the temperature hascompletely stabilised at 60-61°.
3The molds I use are from Polysciences, and are called 'Peel-A-Way' (Cat. no. 18986).

36

In situ hybridization to embryo sections using 35S riboprobes

bunsen flame, bring it to the temperature of the wax sitting in the water bath, andthen pipette the embryos. You want to put in enough wax/embryos to cover thebase of the mold in a thin layer. The number of embryos should be enough to form amonolayer in the mold: fewer will mean that there are fewer embryos in each section;
more will stop the embryos from settling horizontally in the wax.
7) Allow the embryos to settle for about one minute, and then try to make them moveinto the centre of the mold. I do this by blowing the surface of the wax with a pasteur
pipette, and thus pushing the embryos away from the sides. This takes a little time toperfect - it does not really matter if the embryos are spread around the whole base.
8) Fill the mold with wax by slowly and gently, running more wax down the side; try
not to disturb the embryos. The wax should fill the mold to about two thirds of its
volume. Don't let the wax solidify before adding more, as this will form a weakregion, and the block may break.
9) Carefully remove the mold from the hotplate and allow it to set undisturbed for about
15 minutes.
10) The blocks can now be stored at 4° for several months before cutting sections.
Subbing slides:
1) The microscope slides are subbed in poly-lysine to make the the sections stick to themwell. The slides can be subbed in large batches, and stored in a dust-freeenvironment.
2) Wash the slides by soaking them in a large container of hot water with somedetergent. Gently agitate them. Rinse thoroughly (e.g. running water for 30
minutes), and finally rinse in distilled water.
3) Load the slides into racks, and dry in an oven.
4) Soak each rack of slides in a solution of 50µg/ml poly-D-Lysine4 in 10mM Tris pH8
for about 10 minutes.
5) Air dry in a dust-free place; this can take 24 hours.
6) Store the slides in boxes at room temperature.
Cutting sections:1) The exact procedure used will depend on the microtome that you use. The following
notes should provide a guideline.
2) Push the wax block out of the mold, and mount it onto a wooden microtome block,using molten Paraplast Plus to stick it.
3) Trim the block with a razor blade so that the face to be cut is a trapezoid shape.
4) Mount the block onto the microtome, and cut 6 micron sections. The number that you
can fit onto each slide depends on the size of the sections and the size of the coverslip
that you intend to use for the hybridisations. I use 22mm2 coverslips, and use about
4I use Sigma #P-0899, which has a molecular weight range of 70-150,000.

37

In situ hybridization to embryo sections using 35S riboprobes

four sections per slide. I usually cut individual sections rather than ribbons, but bothare OK.
5) Place a subbed slide onto a hot-plate set at 45°. Put a drop of clean water onto theslide, and carefully place the section onto the water drop5. It is important to put the
newly cut (shiny) side of the section downwards. The section will spread flat over
the water, and will be lowered slowly on to the slide as the water evaporates. Afteryou have put enough sections onto a given slide, leave it on the hot-plate,undisturbed.
6) Leave the slides on the hot-plate overnight (or in a 45° oven).
7) The sections can be stored at room temperature in a dry box for several weeks prior to
hybridisation.
Making the 35S riboprobe:
1) The template DNA should be linearised downstream of the insert. In order to remove
any contaminating RNAses, treat the digest with 200µg/ml proteinase K for 30
minutes at 37°, phenol extract once, and chloroform extract. Ethanol precipitate andresuspend at 1µg/µl.
2) I use the Stratagene transcription kit, and follow their protocol:
1µg DNA1µl 10mM rATP
1µl 10mM rCTP
1µl 10mM rGTP
1µl RNAse Block6
1µl 0.75M DTT
4µl 5X transcription buffer (Stratagene)
5µl 35S UTP 40µCi/µl, 800Ci/mmole (SP6/T7 grade)
1µl of a 1:5 dilution of T3 or T7 polymerase in 1X transcription buffer
Final volume = 20µl
3) Incubate at 30° for 1 hour.
4) Add 10X MS to 1X, and 1µl of DNAse (use RNAse-free DNAse). Incubate at 37° for 30
minutes.
5) TCA precipitate a tiny aliquot, to discover the percentage incorporation of label,
which allows to you calculate the amount of probe transcribed.
6) Add tRNA (phenol extracted, ethanol precipitated, and resuspended in DEPC treatedwater). Since you are transcribing new RNA, the total amount made depends on theamount of the hot (limiting) nucleotide in the reaction, and the incorporation. To get
the right probe and tRNA concentration for the hybridisation, use the following
example as a guide. For 75% incorporation, using 200µCi of UTP in the reaction, add
500µg tRNA. For different amounts of transcripts, add an amount that will give the5A small paintbrush is an easy way to handle the sections.
6Stratagene's RNAsin

38

In situ hybridization to embryo sections using 35S riboprobes

same relative amount (i.e. for 50% incorporation of the same amount of label, add
333µg).
7) Phenol extract once; back extract with an equal volume of 10mM DTT; chloroform
extract once; add an equal volume of 4M ammonium acetate (DEPC treated), and 2.5
volumes of ethanol.
8) Chill, spin, rinse pellet, and resuspend in 50 µl of 10mM DTT.
9) Add 50µl 2X carbonate buffer, and incubate at 60° for 140 minutes7.
10) Add equal volume of 0.2M sodium acetate, 1% acetic acid; add 2.5 volumes of
ethanol, chill, spin, rinse pellet, and resuspend in 50% formamide8. Aim for 5 x 105
cpm/µl in the formamide. (For the example above - 75% incorporation of 200µCi - use
96µl of formamide).
-The probe is now 5X concentrated. Store it at -20° till use.
Prehybridisation Treatment of the Slides:
1) Load slides into a rack.
2) The following treatments should be carried out in glass dishes: the dishes I use hold400ml.
3) Dewax in xylenes, twice, for 10 minutes each.
4) Rehydrate in ethanol series: 100%, 95%, 80%, 60%, 30%, 2 minutes each.
5) Incubate in 0.2M HCl for 20 minutes at room temperature.
6) Rinse in H20, 5 minutes.
7) Incubate in 2X SSC, 30 minutes, 70°.
8) Rinse in H2O, 5 minutes.
9) Digest with 0.125mg/ml protease in P buffer at room temperature for 10 minutes9.
10) Incubate in 0.2% glycine in PBS for 1 minute.
11) Rinse in PBS, twice, 1 minute each.
7This is what I've found best to give transcripts with a mass average of around 100 bases. It is

wise to run a small aliquot on a sequencing gel to check the size. Aim for no smaller than 50
bases, and no larger than 200 bases (average size).
8The formamide used for resuspending the probe, and that used in the hybridisation buffer,should probably be of the best quality available. It should be deionised, and perhaps evenrecrystallised. The formamide used in the wash buffer can come straight out of the bottle.
9 I use Sigma protease (#P-5147). It should be made up at 40mg/ml in water, and predigested at
37° for 4 hours. It is then stored in 1ml aliquots. It may need to be titrated: I use the above
conditions, but if these seem incorrect, use half the amount that completely removes the sections
from the slides.

39

In situ hybridization to embryo sections using 35S riboprobes

12) Post-fix in 4% paraformaldehyde in PBS10, 20 minutes, room temperature.
13) Rinse once in PBS, 1 minute.
14) Acetylate the sections in 0.5% acetic anhydride in 0.1M triethanolamine, pH8.0 (I use
3ml in 600ml), 10 minutes, room temperature, in the hood, with vigorous stirring.
Acetic anhydride is very unstable in water - add it to the triethanolamine at the sametime as the slides.
15) Rinse in PBS, 2 minutes.
16) Dehydrate in ethanol series: 30%, 60%, 80%, 95%, 100%, two minutes each.
17) Air dry.
-The hybridisations should be set up soon after this pretreatment - the sections areprobably not very stable at this stage.
Hybridisation:
1) The probe concentration during the hybridisation should be around 100,000cpm/µl.
2) Boil the probe (which is in 50% formamide, 5mg/ml tRNA) for 2 minutes (3µl foreach slide to be hybridised).
3) Chill on ice and add 4 volumes of hybridisation buffer. Mix well, and spin for 30
seconds.
4) Place 15µl on to each slide, near the edge of the tissue sections.
5) Carefully lower a 22mm2 clean (but not necessarily siliconised) coverslip over theliquid in such a way as to make the probe cover the sections. Try not to allow too
many bubbles.
6) Seal the edge of the coverslip with rubber cement; use plenty. I find it easiest to poursome into a 5ml syringe and use it from the syringe.
7) Hybridise overnight in a humid chamber at 50° (not neccessary, but not a bad idea).
Washing:
1) Carefully peel off the rubber cement, using a sharp pair of forceps; try not to dislodge
the coverslip while doing this.
2) Place the slide in a rack in a tank of wash buffer at 50°. Do this, and all subsequent
washes in the hood - the wash buffer is very smelly. Suspend the rack so that there is

10This must be made fresh. Heat it until near boiling, and then add a drop or two of 1M NaOH.

It should go into solution within a few minutes.

40

In situ hybridization to embryo sections using 35S riboprobes

space for the coverslips to slide off, which should happen within a few minutes. Ifneeded, stir the wash buffer gently11.
3) When the coverslips are off, move the rack into a fresh tank of wash buffer. Wash forabout 4 hours at 50°, with hourly changes of buffer.
4) Wash the slides in 1X NTE at 37°, for 5 minutes.
5) Incubate in 20µg/ml RNAse A (the stock solution is 10mg/ml, boiled for 5 minutes,
allowed to cool slowly, and stored at -20°) in NTE at 37° for 30 minutes. This step
removes single stranded RNA, and reduces background.
6) Wash in NTE at 37° for one hour with 4 changes.
7) Dehydrate through ethanol series: 30%, 60%, 80%, 95%, 100%. Dilute the ethanol with
0.3M ammonium acetate instead of water.
8) Air dry.
Autoradiography:
1) I use Kodak NTB2 emulsion, diluted 1:1 with water12, and aliquotted into 5ml
scintillation vials, which are stored in absolutely light tight conditions at 4°13. This is
the right quantity to fill the dipping chamber that I use, which is made of perspex andis just larger than a slide: any vessel that is deep enough to allow the slide to bedipped so that the emulsion easily covers the sections is suitable, but adjust thealiquot size accordingly.
2) Set up a water bath at 45° in the dark room and melt an aliquot of emulsion. I put thevial inside a film developing tank which has some water in it, and put the wholething in the water bath. This is then light tight, and you can come and go safely.
Alternatively a dark room with a properly sealed rotating door can be used. The
emulsion takes about 15 minutes to melt.
3) While dipping the slides and drying them, they must be kept out of all light otherthan weak safelight of the appropriate kind for the emulsion used (Kodak no. 2 for
NTB2). The way that you organise the dark room depends on your set up, just makesure that it is really light-tight.
4) Pour the melted emulsion into the dipping chamber, which should be sitting in thewater bath, to keep it warm, and thus to stop the emulsion from solidifying. Dip each
11The wash buffer is 50% formamide, and therefore should not be put down the sink.

Furthermore, the first couple of washes contain significant amounts of radioactivity, and shouldbe treated with appropriate care. I bottle everything and leave it to EH&S.
12The new batch of emulsion is melted in the dark at 45° (which takes about 45 minutes) andmixed in a conical flask with an equal volume of distilled water, also prewarmed to 45°. The
mixing must be done gently, and bubbles should not be allowed to form: vigorous stirring
increases the background. It is worth checking the background of a new batch, by dipping aclean slide and developing it. Very few silver grains should be visible. Also this same control
can be done with an experiment, which helps to trace the source of any high background.
13Also make sure that there is no radioactivity stored in the same fridge, as the emulsion is very
sensitive, and will develop a high background.

41

In situ hybridization to embryo sections using 35S riboprobes

slide briefly into the dipping chamber and remove it without rubbing off the layer ofliquid emulsion. Place the slide into a rack so that it stands vertically with the end
that you held while dipping, downwards. This allows excess emulsion to drain awayfrom the sections, and causes there to be a thin, uniform layer over them.
5) Allow the slides to dry for at least two hours before putting them in light tight boxes,
which should have some drying gel (e.g.. Drierite) in them. Tape the boxes shut, andwrap them extensively in aluminium foil. Expose at 4°, away from any radioactivity.
6) It is convenient to have three or four slides for each probe used, and to put one of eachtype into each box. This way when you want to develop them, you need only open
one box, and you need not re-wrap it.
7) Try developing the first set of slides after about 4 or 5 days - moderate to strong
signals should be exposed enough by then. Judge how much longer to leave theother slides from the amount of signal seen at this time.
Developing the slides:
1) It is crucial that all the solutions used in the developing and staining of the slides areat the same temperature, which should be below 20°. If they are not, the emulsion,
which is very delicate when wet, may come off the slides during their treatment,
thereby ruining the whole experiment. To avoid this I make up the solutions the day
before using them, and let them all equilibrate in an 18° room. When they are to be
used they can then be taken into the dark room, and it is OK if they all warm up
slowly together. The solutions are:Kodak D-19 developer
2% acetic acid
1:3 dilution of Kodafix fixer14
3 litres distilled water
10mM sodium phosphate buffer, pH6.8
2) Allow the box of slides to come to room temperature while still sealed (about 1 hour).
3) In a dark room with appropriate safelight, load the slides into a rack.
4) Immerse them in developer for 2 minutes.
5) Transfer them to 2% acetic acid for 30 seconds (this acts to stop the developer, and isprobably not crucial).
6) Incubate the slides in fix for 5 minutes.
7) Rinse them in distilled water for 15 minutes with two changes. They are safe in thelight from this point onwards.
8) The final process is to stain the slides with Giemsa. The degree of staining wanted
varies with the intensity of the signal: if the signal is strong, stain for about 15 or 20
minutes, but if it is weak, stain for only 30 seconds to 1 minute. I usually stain thefirst batch to be developed weakly, and then decide how to treat the next batch.
14The type of fixer is not crucial, but is should have a hardener in it. Use the dilution suggested

for films rather than paper.

42

In situ hybridization to embryo sections using 35S riboprobes

9) Make up a 5% Giemsa solution in 10mM NaPO4 immediately before staining theslides.
10) Incubate the slides in the Giemsa solution for the desired amount of time.
11) To remove the stain, gently pour water into the container until the film that forms at
the surface of the Giemsa solution is poured away. This is important so that the film
does not coat your slides.
12) Rinse the slides two or three times in water for about 30 seconds. While wet, they
can be viewed at low power, and if they appear overstained, they can be rinsed more
extensively: normally this is unnecessary.
13) Air dry the slides, and then mount them in DPX mounting medium under acoverslip.

43

In situ hybridization to embryo sections using 35S riboprobes

Solutions:
P buffer:
50mM Tris 7.5, 5mM EDTA.
5X Transcription Buffer:
200mM TrisHCl, pH8, 40mM MgCl2, 10mM spermidine, 250mM NaCl.
10X MS:
100mM Tris pH7.5, 100mM MgCl2, 500mM NaCl.
2X Carbonate Buffer:
80mM NaHCO3, 120mM Na2CO3, which should be pH10.2.
Hybridisation Buffer:
The hybridisation mix can be made in a large volume, and stored at -20°15. The recipe
for 1.25X buffer is:
50% formamide
12.5% dextran sulphate
0.375M NaCl
12.5mM Tris pH7.5
12.5mM sodium phosphate buffer pH6.8
6.25mM EDTA
1.25X Denhardt's
12.5mM DTT
Wash buffer:
1X salts, 50% formamide, 14mM mercaptoethanol.
10X salts:
3M NaCl, 0.1M Tris, 0.1M NaPO4, 50mM EDTA, pH6.8.
Quantities for 1 litre:
175.3g NaCl
14.04g Tris HCl
1.34g Tris base
6.78g NaH2PO4.H2O
7.1g Na2HPO4100ml 0.5M EDTA
10X NTE:
5M NaCl, 100mM Tris pH7.5, 10mM EDTA.
References-Cox et al., Dev Biol, 101, 485 - 502 (1984)-Ingham et al., Nature 318, 439 - 445 (1985)

15The solutions used to make this buffer should probably be DEPC treated, and the whole thingshould be filtered through a millipore filter (which will be slow, because of the dextran sulphate).

44Marek Mlodzik & Nipam Patel

WHOLE MOUNT IN SITU HYBRIDIZATION WITH A NONRADIOACTIVE
PROBE
This protocol describes a procedure for the localization of RNA transcripts
within whole Drosophila embryos using a non-radioactive detection method. The
embryo hybridization procedure is essentially as described by Tautz and Pfeifle. Mostof the modifications are in the procedure for the production of the probe. We have
employed two techniques: a modified version of the standard oligo priming procedure
and a protocol using PCR that generates single-stranded DNA probes.
Recently, Boehringer Mannheim released digoxigenin-UTP for the production of
RNA probes. This could increase the sensitivity of the detection and replace theproduction of DNA probes. However, since this method has not been used yet, it is notincluded in this protocol. The pretreatment and detection as described here are the samefor DNA or RNA probes.
Preparation of Whole Mount Embryos:
This protocol is a slightly modified version of the protocol of Tautz and Pfeifle
which is based on the fixation protocols of Ken Howard, Phil Ingham and AlfonsoMartinez-Arias and the whole mount hybridization protocol of Paul Mahoney and the
Boehringer Genius Kit.
1) Collect the embryos (anywhere between 0-15 hrs) and rinse thoroughly with water.
2) Dechorionate in 50% "Clorox" (commercial bleach) for 2-5 min.
3) Wash with water.
4) Transfer the embryos (up to 3ml volume of embryos) into a polypropylene tube
containing:
16 ml
0.1 M Hepes, pH 6.9
2 mM Mg-sulfate
1 mM EGTA
4 ml
20% paraformaldehyde (dissolve
paraformaldehyde in boiling water.
You may have to add NaOH to get it
into solution)
20 ml
heptane5) Place on a rotator for 15 to 20 min.
6) Remove the lower phase (fixative) and add 15 ml methanol and shake the tube
vigorously for 10-30 sec. The devitellinized embryos will sink to the bottom (heptane
will be on top and the vitelline membranes will stay at the interface).
7) Transfer embryos together with some methanol into a new tube. Wash with
MeOH:EGTA (90% methanol;10% 0.5 M EGTA, pH 8.0).
8) Prepare 4% paraformaldehyde in PBS (PP; made by diluting 20% paraformaldehyde
solution with 1X PBS) and MeOH:EGTA (ME) solution (see above). Rehydrate theembryos as follows:
5 min
in 7 parts ME / 3 parts PP

45

Whole mount in situ hybridization with a nonradioactive probe

5 min
in 5 parts ME / 5 parts PP
5 min
in 3 parts ME / 7 parts PP
20 min in PP9) If you want to store the embryos at this point, wash them in PBS and dehydrate themthrough 30%, 50% and 70% ethanol and store at -20°C; rehydrate before
pretreatment.
II. Pretreatment:
All the following incubation steps are performed at room temperature. Avoid
potential RNAase contaminations. The PBS can be treated with DEPC and autoclaved
and then supplemented with 0.1% Tween 20 (=PTw). We have found that the DEPC
treatment is not absolutely necessary.
1) Wash the embryos 3 times for 5 min each in PTw
2) Incubate the embryos for 7-8 min in PTw plus 50 µg/ml non-predigested Proteinase
K (Boehringer Cat# 745 723; you may have to determine the optimal extent oftreatment for your batch of proteinase K; the optimal time seems to be 2-3 min before
the embryos begin to show any noticable damage).
3) Stop Proteinase K digestion by incubating for 2 min in 2 mg/ml glycine in PTw.
According to Tautz andf Pfeifle, the Proteinase K digestion is a critical step. If it
is too short, you lose sensitivity and increase the background. If it is too long, theembryos tend to disintegrate during hybridization.
4) Wash 2 times for 5 min each with PTw.
5) Refix for 20 min with PP.
6) Wash 5 times for 5 min each in PTw.
Preparation of DNA Probe
I. PCR-labeled Single-stranded Probe from Cloned DNA:
Using this PCR technique, single stranded DNA probes containing digoxigenin
can be made for in situ hybridization. The biggest advantage of this technique over the
random oligo priming method is that a much larger quantity of probe can be made withthe same amount of starting nucleotide. In addition, the ratio of labelled DNA to
unlabelled starting material (which will compete for hybridization) is much higher. In
situations where transcripts are not very abundant, these single-stranded probes seem towork better than the oligo primed probes. The major disadvantage is that it is difficult
to control the probe size. The PCR reaction will generate full length strands for inserts
up to 2-3 kb. We simply boil the DNA to reduce its size, but have not made any
estimates of how effective this really is nor have we investigated other alternatives tothis problem. Empirically, however, this labelling technique works extremely well forthe in situs. In addition, we have found that biotin-16-dUTP can also be incorporated
this way and used for in situs using streptavidin-alkaline phosphatase for the detection
step. This method works okay, but is clearly 2 to 3- fold less sensitive than thedigoxigenin. Also, PCR can be used to generate very large amounts of biotin-labelleddouble-stranded DNA by the use of two primers as long as the distance between
primers is less than 2-3 kb. Finally, we have observed that biotin-labelled DNA runs

46

Whole mount in situ hybridization with a nonradioactive probe

normally in agarose gels, but digoxigenin-labelled DNA will run as though it is larger
than it actually is.
1) Prepare the following stock solutions:
10 x React
0.5 M KCl
(comes in PCR kit)
0.1 M Tris-HCl, pH 8.3
15 mM MgCl2
0.01% (w/v) gelatin
5 x dNTP Mix 1 mM dATP
7.1 µl of 10mM
(equivalent to 1 mM dCTP
7.1 µl of 10mM
tube 6 of Genius
1 mM dGTP
7.1 µl of 10mM
kit)
0.65 mM dTTP
4.6 µl of 10mM
0.35 mM Digoxigenin-11-dUTP
25.0 µl of 1mM(Dig-dUTP can be purchased 20.5 µl of H2O
as a 1mM soln from Boerhinger)
71.4 µl total vol.
SK primer
KS primer

30 ng/µl (approx 5.3 µM)
30 ng/µl (approx 5.3 µM)

Linearized DNA containing insert. Linearize Bluescript DNA as you would to makean RNA runoff probe. For example, you might have a 2.0 kb EcoR1 fragment cloned
into the R1 site of Bluescript. You might linearize by cutting one aliquot with Kpn
and another with SacII (assuming there are no Kpn or SacII sites in the insert). The
Kpn cut DNA can be used with the SK primer to create one strand which might be theanti-sense strand. The SacII cut DNA can be used with the KS primer to create thesense strand which can be used as a control in the hybridization. T3, T7, M13, orinternal primers can be used in place of the KS/SK primers. We have on occasion cut
DNA and then not bothered to phenol extract and precipitate afterwards, but have
simply heat inactivated the restriction enzyme and then successfully used it for the
PCR reaction. You will want to dilute your DNA with water to a final concentrationof about 100-200 ng/µl. Keep in mind that if your insert is much over 3 kb that theprobe produced will probably not be fully representative of the entire insert .

47

Whole mount in situ hybridization with a nonradioactive probe

2) Set up the following reaction (25 µl volume):water
8.5 µl10 x React
2.5 µl5 x dNTP mix
5.0 µlKS (or SK) primer
5.0 µlDNA (100-200 ng/µl) 2.0 µl
Add 40 µl mineral oil and centrifuge. Boil 5 min then add Taq:
Taq I DNA Polymerase
2.0 µl(1.25 units total; a 1:8 dilution in
25.0 µl water of 5U/µl Taq stock)3) Mix the contents and then centrifuge for 2 min. Incubate for 30 cycles in the PCR
thermal cycler under the following conditions:
95°C for 45 seconds
55°C for 30 seconds
72°C for 1 minute4) After the PCR run, add 75 µl dH20, then centrifuge.
5) Remove 90 - 95 µl of the reaction from beneath the oil.
6) Do two ethanol precipitations as follows. Add NaCl to 0.1 M and 3 vols of 100%
EtOH. You can use 10 µg of glycogen or tRNA as a carrier (0.5 µl of a 20 mg/ml
stock). Mix well, -70oC for 30 min, and centrifuge. Wash with 70% ethanol.
Speedvac dry. 2nd precipitation is optional.
7) Resuspend pellet in 300µl of hyb buffer (hyb. buffer of Tautz and Pfeifle protocol
below). For efficient penetration and hybridization to the embryos, the average probe
length should be about 50-200 bp. To reduce the size of the single-stranded DNA,boil the probe for 40-60 min. The probe can be diluted as much as ten-fold before use,
but this varies quite a bit depending on the abundance of your transcript and how
much of a problem you have with background staining. I recommend that you use
the probe anywhere from undiluted (original 300µl) to three-fold diluted for your firstattempt.
8) Check of probe (solutions are same as those described in the embryo protocol below):
1. Remove 1 µl of probe. Add 5 µl of 5X SSC. Boil 5 min. Quick cool on
ice.Centrifuge.
2. Spot 1-2µl onto a small nitrocellulose strip that will fit into an eppendorf tube
or 5 ml snap cap tube.
3. Bake between two sheets of Whatman paper in an 80°C vacuum oven for 30
min. The residual formamide may cause the nitrocellulose to warp. If this is aproblem, reduce the time in the baking oven or do this spot test before thesecond precipitation. Note: unincorporated nucleotide binds only slightly tothe nitrocellulose.
4. Wet the filter with 2X SSC, wash 2X 5 min in PBT. Place strip into eppendorf
tube or 5 ml snap cap tube.
5. Block 30 min in PBT. Incubate in PBT + antibody (1:2000) for 30-60 min.

48

Whole mount in situ hybridization with a nonradioactive probe

6. Wash 4 x 15 min in PBT.
7. Wash 2 x 5 min in NaCl/MgCl2/Tris/Tween solution (Levamisole is notneeded).
8. Develop with NBT and X-phosphate as described for embryos. Spots should
be visible within a few minutes and dark by 10-15 min.
II. Preparation of probe by oligolabelling
An alternative to PCR-generated probes is to create probes by the random oligopriming technique. This is the modification by Charlie Oh (Kornberg lab) of the
protocol that is described in the Genius Kit. By increasing the random primer
concentration and lowering the temperature, smaller probe fragments are createdresulting in a higher signal and less background.
10X LB475 µl 1M Pipes pH 6.6
25 µl 1M MgCl2
3.3 µl ßME
Labelling reaction:
1) Prepare 200ng of purified insert DNA in 9 µl of water.
2) Boil 10 min.
3) Quick freeze in dry ice/ethanol or liquid nitrogen.
4) Add following components on ice.
10X LB2.0 µl
hexamer
6.0 µl of a 10 mg/ml soln. (Random hexamers;
Pharmacia # 27-2166-01)
dNTP
2.0 µl (tube 6 of Genius Kit)
Klenow
1.0 µl (5 Units)
5) Incubate at 15°C for 1 hr.
6) Transfer to room temp for 3 hrs.
7) Precipitate probe and resuspend in 200µl of hyb buffer.

49

Whole mount in situ hybridization with a nonradioactive probe

Hybridization and detection:
1) Hyb solution:

50% formamide
5x SSC
100 µg/ml sonicated salmon sperm DNA
50 µg/ml heparin
0.1% Tween 20

2) Wash embryos 10 min in a 1:1 mix of hyb soln. and PTw.
3) Wash embryos 10 min in hyb soln.
4) Prehybridize in heat denatured hyb soln for about 1 h at 45°C. We have found thatembryos can be stored in the hyb soln at -20°C for 1-2 weeks.
5) Boil the probe for 10 min and then cool on ice. Remove most of the prehyb soln fromthe embryos and add 100-200µl of probe. About a 30µl volume of embryos in the 100200 µl of probe works well.
6) Mix and hybridize overnight in a 45°C waterbath. Round bottom Nunc tubes work
well for this. After the hybridization, the probe can be recovered and used
repeatedly. We have been able to use the same bit of probe up to six times. The
repeated boiling may improve the signal by further reducing the size of the probe.
7) Wash embryos 20 min in Hyb soln. All washes in steps 7 and 8 should be done withagitation at 45oC. You will want to pre-mix and pre-warm all the solutions.
8) Wash embryos 20 min each in:

4 parts Hyb. soln. / 1 part PTw
3 parts Hyb. soln. / 2 parts PTw
2 parts Hyb. soln. / 3 parts PTw
1 part Hyb. soln. / 4 parts PTw
2 times 20 min in PTw

9) Wash embryos 2 times 20 min each with PBT. This and all subsequent steps are doneat room temp.
PBT = 1X PBS, 0.1% BSA, 0.2% Triton X-100.
10) Dilute the alkaline phosphatase conjugated goat anti-digoxigenin antibody (vial 8 of
the Genius Kit) to 1:2000 - 1:2500 with PBT. Incubate the embryos for 1 hour with
200-400µl of the dilute antibody soln.
11) Wash 4 times 20 min each in PBT.

50

Whole mount in situ hybridization with a nonradioactive probe

12) Wash 3 times 5 min each in

100 mM NaCl
50 mM MgCl2
100 mM Tris, pH 9.5
0.1% Tween 20
1 mM Levamisol (Sigma #L-9756; add
fresh just before use.
Levamisole is an inhibitor of
potential endogenous
phosphatase. Nipam has done
experiments where it has beenleft out and there does not seemto be any background problem.)

Prepare a solution of NaCl/MgCl2/Tris/Tween/Levamisole containing 4.5 µl of
NBT (vial 9 of Genius kit) and 3.5 µl of X-phosphate (vial 10) per ml of solution. Add
0.5-1.0 ml to the embryos after the last wash. You will see the signal appear
anywhere between 10-60 min. Stop by washing several times in PBS. We have triedalternative substrates. Vector Kit II substrate produces a brown/black product thatis insoluble in xylene and can therefore be mounted in permount. This substratediffuses less and allows for the resolution of sub-cellular localization, but it is 2-3
fold less sensitive than the standard method. We have not had any luck using an
HRP enzyme detection system.
13) The embryos reacted with NBT/X-phosphate should be transfered to 50% glycerol in
PBS for 1-2 hrs and then transfered to 70% glycerol in PBS. They can be stored at 4°C
for several weeks.
Some Final Notes:
If the embryos are prepared properly, the morphology can be excellent. The
embryos can be dissected in the 70% glycerol soln if needed. One additional possibility
is to reduce the level of background by digesting with a single-stranded nuclease after
the hybridization with a single-stranded probe. This would be equivalent to the RNAse
A digestion of 35S RNA in situs. This should destroy the unbound probe and leave the
RNA/DNA hybrids intact. It may be tricky as the single-stranded nucleases can bedifficult to control. The protocols described above are based on experiments performedby me, Nipam Patel, Yash Hiromi and Elinor Fanning using a variety of different geneprobes.
REAGENTS:
Genius Kit. Boehringer Cat# 1093 657. Alternatively, the components can be purchased
individually. Several are listed below.
Digoxigenin-11-dUTP Boehringer Cat# 1093-088, 25 nmol/25 µl (1mM)
Anti-Digoxigenin-AP Conjugate Boehringer Cat# 1093-274
NBT is 4-Nitro blue tetrazolium chloride, Boehringer, Cat# 1087-479
X-phosphate is 5-Bromo-4-chloro-3-indolyl-phosphate, Boehringer, Cat# 760-994

51

Marek Mlodzik

FIXATION AND PRETREATMENT FOR IMAGINAL DISC WHOLE
MOUNT NONRADIOACTIVE IN SITU HYBRIDIZATION
The following protocol is based on the method described for whole mount embryo
hybridization (see above).
1) Dissect discs from late 3rd instar larvae in PBS.
2) Fix dissected material for 15'-20' in 4% paraformaldehyde, PBS, 0,5% Triton X-100 on
ice in Eppendorf-tube, while you continue to dissect more discs.
3) Fix for additioinal 20' at RT (same fixative as above) after you have finisheddissecting.
4) Following fixation, wash the discs 3 times for 5' in PTw at RT.
5) ProteinaseK digestion:
12.5 µg/ml enzyme in PTw (1/4 of the concentration that is used for embryos)
usually between 4' and 10' at RT is fine, depends on batch of proteinaseK
when new batch: titrate time !
The longest digestion that still retains reasonable morphology gives the best
results.
6) Wash digested discs in PTw, 2mg/ml glycine: 2 times 5' at RT, followed by two 5'
washes in PTw.
7) Postfixation: 20' at RT in

4% paraformaldehyde
0.2% glutaraldehyde
PTw8) Wash fixed discs in PTw 4 times 5' at RT.

Prehybridization:1) PTw:

hybridization buffer (1:1) for 10' at RT

2) Hybridization buffer, 10' at RT
3) Hybridization buffer, 1 hour at 45 C
4) Hybridize o/n at 45 C, in Eppendorf tube without shaking

52

Fixation and pretreatment for imaginal disc whole mount nonradioactive in situ hybridization

Buffers:
PBS (10X stock)76.1 g NaCl
18.8 g Na2HPO4.7H2O (9.9 g if anhydrous)
4.1 g NaH2PO4.H2O
H2O to 1 litre

PTw

PBS containing 0.1% Tween

Hybridization Buffer same as for embryos (see above)
Probe and Detection:
Probe can be made either by random-oligo priming or alternatively by PCR generating
single stranded labelled DNA (see above).
More recently, a digoxigenin-labelling kit for RNA probes is also available.
Detection is done as described for embryos (see above), all steps in Eppendorf tubes.
Comments:
The success of the method depends largely on the abundance of the respective
transcripts. In our experience, detection of transcripts that are found at 1:50,000 (ormore abundant) in the eye disc cDNA library usually work fine, e.g. glass (1:5000) or
scabrous (1:20,000). For transcripts with lower abundance, e.g. seven-up (1:100,000) or
sevenless (1:100,000), this method works in about 1 out of 2 experiments.
GOOD LUCK !

53

Matthew Freeman

IMMUNOSTAINING DROSOPHILA EMBRYOS
Fixation and permeabilisation.
1) Collect embryos on grape juice agar collection plates, and rinse them onto a Nitex
mesh sieve, rinsing them well with water containing 0.05% Triton-X-100 (optional) to
remove excess yeast.
2) Dechorionate in 50% household bleach (2.5% sodium hypochlorite final conc.), by
dipping the whole Nitex sieve into a beaker of bleach for 2-4 minutes and swirling theembryos once or twice. Rinse well in PBS/0.05% Triton-X-100 to remove the bleach.
Do not let the embryos dry out at this stage.
3) Transfer the embryos with a soft paintbrush to a vessel with a tight-fitting lid
containing:
50% n-Heptane
50% PEM-formaldehyde
PEM-formaldehyde is:
0.1M PIPES
2mM EGTA
1mM MgSO4
pH to 6.95 with HClmix 9:1 with stock solution of 37% formaldehyde
The size of the vessel, and the quantity of fixative should be adjusted according
to the number of embryos used. For about 50-500 emryos I use a 5ml glass vial with ascrew-cap; for 0.5-1g of embryos I use a 50ml polypropylene tube with a screw cap. It
is a good idea to use enough fix/heptane to fill the container to at least 70-80%, as thisstops too many embryos being lost by sticking to the sides.
This fix is a good all-purpose one, but there may be some antigens which aredestroyed by it. The most likely source of problems is the formaldehyde solution,
which contains up to 10% methanol as a stabiliser. Some antigens (for example 24B10
antigen) are methanol sensitive, in which case, either buy expensive and unstableformaldehyde without methanol, or make it by dissolving paraformaldehyde inwater. To do this heat the water to near boiling and add a drop or two of 1M NaOH.
You can substitute PBS or Balanced Salt Solution (for recipe see Wilcox, in Roberts:
Drosophila, A Practical Approach. IRL Press. 1986) for the PEM, but I like PEM thebest for most cases. Other fixes are possible: anything you know preserves yourantigen should work, though there might be penetration problems if you use too higha glutaraldehyde concentration.
4) Vigorously shake the embryo/heptane/fix for 15-30 minutes at room temperature.
5) With a pipette, remove as much of the aqueous (bottom) layer as possible, without
taking the embryos at the interface.
6) Add 1 volume of 100% methanol to the embryos/heptane, and shake very hard for 30
seconds-1 minute. Allow the two phases to separate. The devitellinised embryos willsink to the bottom, and the embryos with their vitelline membranes still attached andthe empty vitellines will remain at the interface. This step is not always very efficient:
anything over 50% sinking is fine, and on a good day it should be 80 or 90%.

54

Immunostaining Drosophila Embryos

7) Remove the heptane (top layer), all the interface, and the methanol above theembryos with an aspirator.
8) All the subsequent steps are carried out in the same tube, which can be filled with asolution, the embryos allowed to settle, and the solution removed.
9) Treat the embryos with:
100% methanol

10 mins (at this stage the embryos can be
stored at -20°C for months)
PBS/0.2% saponin
10 mins""
10 mins
The embryos can be stored at 4° like this for a week or two.
Before staining:
PBS/0.2% saponin/5% serum (normal goat or foetal calf)
10 mins

As discussed above, some antigens are methanol sensitive, and in this case thevitelline membranes have to be removed by hand. In this case, substitute the followingfor 6-9 above:6a) Remove the aqueous (lower) phase. Take some embryos in heptane with a pipette,and spread them dropwise onto a microscope slide covered with double sided sticky
tape. Let the heptane evaporate, and just as it does, immerse the slide in PBS/0.2%
saponin in a petri dish; do not let the embryos dry out on the slide. Using a finetungsten needle, gently tease the embryos out of their (already cracked) vitelline
membranes; this step is made much easier if the embryos are initially fixed for longer
than normal (about 45 minutes). Transfer the embryos to an Eppendorf tube of
PBS/0.2% saponin/5% serum, using a Gilson pipette with a yellow tip cut off tomake a wide enough opening.
Antibody Staining:
I stain the embryos in Eppendorf tubes or, for large numbers of embryos, 12x75mm
polypropylene tubes (Falcon 2063).
1) Incubate the fixed embryos in primary antibody diluted in PBS/0.2% saponin/5%
serum. Typically monoclonal supernatant is diluted about 1:5, polyclonal serum
about 1:500, and ascites even more, but you should determine the best for yourantibody. Incubate for 2 hours at room temperature, or overnight at 4° (which isprobably a little more sensitive).
2) Rinse in 3 changes of PBS/0.2% saponin for 10 minutes each (longer washing may
reduce background).
3) Incubate in secondary antibody diluted in PBS/0.2% saponin/5% serum for 2 hours toovernight. In most cases, I use peroxidase conjugated 2nd antibody, and do a DAB
reaction; this is the most sensitive detection system available, and can be made evenmore so with silver-gold intensification. However, a fluorescent conjugated 2nd
antibody may be used, in which case, I prefer rhodamine or Texas red to fluoroscein,since they are less prone to quenching under epifluorescent illumination.
4) Rinse in 3 changes of PBS/0.2% saponin for 10 minutes each, and one rinse of PBS. For
fluorescent labelled embyos mount in 85% glycerol containing 2.5% isopropyl gallate,and view with epifluorescent illumination. An optional Hoechst 33258 stain can be
added to the penultimate rinse (1µg/ml; caution - mutagen); this will strongly label

55Immunostaining Drosophila Embryos

all nuclei, and is viewed with ultraviolet epifluorescence. Slides mounted in glycerol
dry out after a few days unless the edge of the coverslip is sealed with nail varnish.
5) For the DAB reaction, when using peroxidase-conjugated 2nd antibody, incubate theembryos in 400µl of 0.5mg/ml diaminobenzidine in PBS. After 10 minutes, add 2µl of
3% hydrogen peroxide and watch the development of the brown reaction productunder a stereomicroscope. When the reaction is sufficient (usually 5-15 minutes), stop
it with 2 or 3 rinses in PBS. (DAB can be made up in 10X aliquots (5mg/ml in water)and stored at -70°. USE GREAT CARE WHEN WORKING WITH DAB. IT MAY
BE A VERY POTENT CARCINOGEN. Use copious amounts of bleach to inactivate
it (leave overnight), and then rinse it away with large amounts of water. Treat gloves,tips, and tubes in the same way.)6) Either take the embryos through silver-gold intensification (see Liposits et al.
Neuroscience 13, 513-525. 1984 and the previous edition of the Rubin Lab Manual) or
mount them in 85% glycerol, or dehydrate them through an ethanol series, clear themin methyl salicylate until they sink, and then mount them in the methyl salicylate(this gives the best results, but is less straightforward than aqueous glycerol).
Note:
In this protocol, I have suggested using 0.2% saponin throughout the staining. This isa good, very mild detergent which should not extract many antigens. However, if it
does not work, you could try using no detergent (which makes the embryos sticky, andreduces the ease of antibody penetration), or substituting another detergent. If yourantigen is nuclear, 0.1% Triton-X-100 might be better.
Reference:
The best overall discusssion of these, and many other, techniques is:
Immunocytochemistry: Theory and Practice. Lars-Inge Larsson. CRC Press (1988)

56

N.E. Baker
STAINING EYE IMAGINAL DISCS
A. Labelling with antibodies:
Labelling eye discs with antibodies does not differ in principle from labelling anyother tissue. However, there are several manipulations that aid treatment of thistissue. The following protocol is from Tomlinson and Ready, Devl. Biol. 120, 366-376(1987). A second section gives detailed comments and tips on some of the steps, withthe aim that the protocol could be followed by an inexperienced worker without
further instruction.
1)

Dissect eye imaginal discs from larvae in 0.1M NaPO4 (pH 7.2). It is convenient
for subsequent handling to leave the discs attached to the mouthparts. (Seedetailed instructions.)

2)

Fix the discs. Usually we use PLP for 45 min on ice. PLP gives fixation resultscomparable to glutaraldehyde but preserves antigenicity. Of course theinteraction of each antibody with its antigen is unique and may require individual
treatment. Less commonly we use 4% paraformaldehyde in PEM for 30 min atroom temperature, particularily with nuclear antigens (PLP is a better membrane
fix and we guess may reduce accessibility of the nucleus to antibodies). (Seedetailed instructions.)

3)

Wash discs in 0.1M NaPO4 (pH7.2), 0.1% saponin for 15 min on ice.

4)

Dissect away the peripodial membrane. Removing the peripodial membraneincreases accessibility of the epithelium to antibody and visibility of the stained
tissue under the microscope. For some antibodies the improvement is neglegible(eg: Mab BP104). For others this step is essential (eg: Mab22C10). (See detailed
instructions.)

5)

Incubate in primary antibody, 1-2 hr on ice. Make up antibody in 0.1M NaPO4(pH7.2), 5% normal goat serum, 0.1% saponin (store frozen or filter sterilized at4°C). Use monoclonal supernatants at 1:1 dilution (but Mab22C10 at 90%), ascites
or sera at 1/250 - 1/5000 as appropriate. Final saponin concentrations of 0.01% 0.1% are satisfactory for light microscopy. (See detailed instructions.)

6)

Wash 3X in NaPO4 + serum + saponin.

7)

Incubate in secondary antibody, 1-2 hr on ice. Generally we use HRP-coupledgoat anti mouse/rabbit from BioRad at 1/200 - 1/500.

8)Wash 3X in NaPO4 + serum + saponin.

9)

Incubate in diaminobenzidine (for HRP coupled secondary). The reaction is
generally complete after 15-30 min, and for many antibodies staining can be muchbriefer. Staining solution is 0.5 mg ml-1 DAB, 0.1% saponin, 0.1 M NaPO4 pH7.2,
0.003% H2O2. (See detailed instructions.)

57

Staining eye imaginal discs

10) Wash in 0.1M NaPO4 .
11) Transfer to a screw-cap vial in 0.1M NaPO4, such as the 1 dram/3.7 ml bottles
from Fisher.
12) Postfix and intensify in 2% OsO4 in 0.1M NaPO4 , from 2min on ice to 10 min atroom temperature, as desired. (See detailed instructions.)
13) Wash in several changes of 0.1M NaPO4. Discs can be stored at 4°C at this stage.
14) Using a pasteur pipet, transfer to 50% ethanol in a petri-dish (60x15mm). After 5
min transfer to 100% ethanol, and then again to fresh 100% ethanol.
15) Mount in DPX.

(See detailed instructions.)

Detailed Instructions for steps numbered:
1) To remove eye-antennal discs, hold the larva firmly about halfway down the body,using forceps. With a second pair of forceps, grasp the mouthparts and pull themout of the head. Usually the eye-antennal discs come out with the mouthparts.
Other tissues may also remain attached; still holding on to the mouthparts,
remove the brain and salivary glands, if present. Then grab the internal part of themouthparts (forceps B in Fig. 1) and remove this and the eye-antennal discs fromthe external part of the mouthparts (still held by forceps A in Fig. 1). All othertissues should get left behind at this step.
To make 1 liter 0.1 M NaPO4 pH7.2 use:
10.22 g Na2HPO4.H2O OR
12.82 g Na2HPO4.2H2O OR
19.30 g Na2HPO4.7H2O OR
25.79 g Na2HPO4.10H2O plus 3.86 g NaH2PO4.H2O OR
4.37 g NaH2PO4.2H2O

FIGURE 1
2) The manner in which the discs are transferred to fixative is important. Flatter
preparations, better for both analysis and photography, will be obtained if discs
are spread on the surface meniscus, apical surface downwards. Use a wire hook
(0.51 mm dia. tungsten wire from Ted Pella) to pick up the tissue where theantennal discs are attached to the mouthparts, and lower it into the fix.

58

Staining eye imaginal discs• PLP is 2% paraformaldehyde, 0.01 M NaIO4, 0.075 M lysine, 0.037 M NaPO4,
pH7.2. For light microscopic work, we do not pH the final solution, but instead
make up PLP as follows. Make an 8% formaldehyde solution fresh from solid
without boiling (a trace of NaOH will put the formaldehyde into solution with
only moderate heating). While this is cooling, dissolve 0.36 g lysine in 10 ml H2O,
7.5 ml 0.1 M NaPO4 pH 7.2, 2.5 ml 0.1 M Na2HPO4 on ice. Immediatley before use,mix 15 ml buffered lysine solution, 5 ml formaldehyde solution, 50 mg NaIO4 togive PLP.
• PEM is 0.1 M PIPES pH7.0, 2 mM MgSO4, 1 mM EGTA, and is conveniently madeup as a 2X stock to be diluted with 8% formaldehyde.
4) We dissect the peripodial membranes on a transparent dissecting dish. One can bemade by setting a thick layer of "Sylgard 184" silicone elastomer (Dow Corning) in
a 150x15mm glass petri dish. Transfer a pair of discs to a large drop of wash
solution on the dissecting dish. Arrange fibre-optic illumination against a black
background so that the peripodial membranes are visible (low-angle illuminationfrom the side opposite you is best).
Good quality forceps and scalpel will help the dissection itself. Use Dumont#5 Biologie grade forceps (expensive, but can be resharpened on a stone whennecessary), and a #11 scalpel blade with agood point (not all brands are the same,e.g. only about 1 in 5 Feather blades is as
sharp as the typical Bard-Parker blade).
To remove the peripodial
membrane, grip the antennal disc toposition the eye disc at the correct angle.
Once the antennal disc has been flattened
it will not be easy to reposition the eye
disc, so first remove other tissue (egpharynx) that may interfere, and run theforcep tips between the two eye-antennal
discs to ensure they can be moved
separately. Rotate the eye-antennal discso the antennal disc is towards you, theeye disc is on edge, and the peripodial
membrane faces right(for a right hander).
Hold the antennal disc firmly to maintainthis orientation (see Fig. 2). Use thescalpel blade to pin the peripodial
membrane to the dissecting dish between
the eye and antennal discs, where it isfurthest separated from the epithelium(see Fig. 2). Using your grip on theantennal disc, pull the eye disc awayfrom the blade (pull down and away,
about 7 o'clock in Fig. 2). Ideally, the torn
peripodial membrane will spring backfrom the eye disc and curl up near the
optic stalk. At other times all or part ofthe membrane remains closely apposedto the epithelium, and it is necessary to
FIGURE 2
peel it off, using forceps to grasp the cut
edge.

59

Staining eye imaginal discs

The apical surface of the epithelium that is now exposed will often adhere to
wire transfer hook, so henceforth always pick up the discs from the basal side.
At first most people find this dissection difficult. General hints that might apply
to any dissection include: 1) Avoid disturbance, stress or stimulants; 2) Brace every
available digit for stability; 3) Put down dissecting implements that are not in use, lest
they dissect something unexpectedly; 4) Know your limits and don't destroy a gooddissection trying to make it perfect. In case a target is useful, with practice themembrane can be dissected and a pair of discs transferred to the next solution in wellunder a minute, on average.
5)

Antibody incubations, washes, and DAB reactions are carried out in the 60-well(conical bottomed) microtest plates manufactured by Nunc. Use a wire hook totransfer the discs from one solution to the next. Each well takes 13 µL.
Incubations can be left overnight at 4°C if the lids are sealed with parafilm to
reduce evaporation.

9)

Unsubstituted benzidene is one of the most potent chemical carcinogens
known. Diaminobenzidene is believed to be safer but should be handled carefully.
Sigma sells DAB in sealed "isopacks" which are convenient for making stock
solution. Aliquots should be stored at -70°C. All materials that contact DAB should
be soaked with bleach prior to disposal (I like to handle DAB inside a heatseal bag,which the waste never leaves).

12)

Osmium intensification produces a background staining of the tissue that we
find useful as a counterstain. Cleaner intensification can be obtained by adding
Co and Ni to the DAB reaction (add 30 µl 50 mM CoCl2 50 mM NiCl2 to 1 ml ofbuffer immediatly before the DAB. Ignore the clouding due to precipitation of
the phosphates)(Adams, J.C., J. Histochem. Cytochem. 29, 775, 1981). In this case a
different postfixation will be required before mounting. 2% glutaraldehyde canbe used (and results in slight destaining). Alternatively a graded series of
ethanols is possible (30%, 50%, 70%, 90%, 100%), but this will not preserve
delicate structures such as the apical photoreceptor-cell membranes. Of course
EM-intensification protocols can also be used, such as the silver-gold method(Liposits et al., Neuroscience , 13, 663, 1984).
SAFETY. Both osmium tetroxide and glutaraldehyde are volatile neurotoxins
with cumulative effects, and should be handled in a fume-hood. Osmium
tetroxide fixes the brain via the nasal mucosa, repeated glutaraldehyde exposure cancause blindness. Osmium tetroxide has a pungent odour, somewhat like garlic.
Glutaraldehyde has only a mild smell. Waste solutions should be inactivated bypouring onto excess "protein" (eg glycine or powdered milk). OsO4 issufficiently volatile that solid waste, eg gloves, is clean if left overnight.

15.

Mount as follows: Take up discs in 100% ethanol using pasteur and drop onto aclean microscope slide. Wipe away excess ethanol with a tissue. Add one dropof DPX mountant (FLUKA) before the discs dry, but when little ethanol remains.
Remove the mouthparts and orient the discs apical side upwards using fineforceps. If desired, remaning peripodial membrane or other debris can beknocked off at this point. Carefully lower a 22x40 mm coverslip down from one
side, watching all the while to see that the discs do not turn over. To some extent
disc rotation can be controlled by moving the coverslip. If this is unsuccessful,
remove the coverslip and reorient the discs. Another drop of DPX will benecessary before replacing the coverslip. As soon as you are satisfied, flatten the60Staining eye imaginal discs

discs with smooth firm pressure on the coverslip (eg: with the butt end of a pairof forceps). Releasen the pressure equally smoothly. Do not attempt to remove
the coverslip once the discs are flattened.
B. Cobalt Sulphide staining:
CoS staining offers a rapid method for visualizing the apical borders of disc cells.
The protocol given here is that of Tomlinson & Ready (Dev. Biol., 120, 366, 1987).
1)

Dissect discs in 0.1 M NaPO4 pH7.2.

2)

Fix in 2% glutaraldehyde in 0.1 M NaPO4 pH7.2 (handle in the fume hood), 30
min, room temperature.

3)

Wash in 0.1 M NaPO4 pH7.2

4)

Wash in H2O.

5)

Dissect away the peripodial membrane as for antibody staining, except that thediscs are still in H2O. Also it is necessary to remove the membrane, not just peel
it back, or else it will snap back over the disc in subsequent steps.

6)

Incubate in 2% CoNO3 in H2O. 96-well microtest plates are useful here too.

7)
8)

Transfer to 1% (NH4)2S in H2O, until the disc is coal-black (typically about 30s)1.
Transfer to H2O. Watch the disc destain. It is about done as the black becomesdark grey and some details can be seen (typically about 30 s). Slightly before thispoint, transfer to a drop of glycerol on a microscope slide and mount under acoverslip. Do not squash the disc, allow it to be gently flattened by the coverslip. The staining is not stable and data should be recorded as soon as possible.
If the coverslip is sealed with nail polish and the slide stored at 4°C the staining
seems to be OK for a few weeks, otherwise 24 hr is about the safe limit.
The stain is essentially a precipitate coating the apical disc surface and isvery delicate. It is easily destroyed if the discs are handled roughly. Once theperipodial membrane has been removed it is important not to handle the apicalsurface during transfers. Make sure to get all the glycerol off the wire hook
before starting each new prep.

C. Basic Fuchsin staining

1Make ammonium sulphide solution fresh from a 20% stock (Fisher Scientific). Keep theammonium sulphide well away from the cobalt nitrate and use a fresh well as soon as any
precipitate forms on the surface of the solutions. Ammonium sulphide gives off H2S gas, which
is more toxic than hydrogen cyanide, but the smell becomes intolerable way below dangerous
levels. Nevertheless, handle the 20% stock in a hood and get worried if you can no longer
smell the stuff.

61

Staining eye imaginal discs

Basic fuchsin is a nuclear stain. Mitotic chromosomes and, at other stages of thecell cycle, spindle pole body associated material are especially well stained. Basicfuchsin is useful for following cell division and cell death (in dying cells the whole cellis labelled). The version used here was adapted from Weischaus and NüssleinVolhard, p 223 in Drosophila, a Practical Approach , (ed D.B. Roberts), IRL,
Oxford/Washington, 1986. Stainless steel mesh baskets, of the kind used forembryonic cuticle preps. are useful for handling the discs.
1)

Dissect discs in 0.1 M NaPO4 pH7.2.

2)

Transfer to Fix A (4 ml 95% EtOH, 0.5 ml CH3COOH, 1.6 ml 20%
paraformaldehyde) at room temperature for 10 min.

3)

Transfer to Fix B (4 ml 95% EtOH, 0.5 ml CH3COOH, 0.4 ml 20%
paraformaldehyde) at room temperature for 1 h.

4)

Wash thoroughly in 70% ethanol.

5)

Incubate in 2 M HCl for 10 min at 60 °C.

6)

Wash with H2O.

7)

Wash with 5% acetic acid.

8)

Stain in 1% basic fuchsin in 2.5% acetic acid for 20 min at room temperature. The
fuchsin solution should be made up fresh every few weeks.

9)

Destain in 5% acetic acid. Do not overdo this step - usually a few minutes isenough. A few salivary glands can be included as a control, as their nuclear
staining is easily visible with the dissecting microscope.

10)

Dehydrate in 70 %, 95% and 100% EtOH, and mount in DPX.

62

Bruce Kimmel

ANTIBODY AND COBALT STAINING OF PUPAL EYE DISCS
Dissection of pupae greater than 40 hrs. old:1) Obtain a bottle or vial of flies which has wandering third instar larvae on the sidesof the vessel. Identify all of the white prepupae in the vessel and circle them with anoneraseable pen noting the time at which each individual was in the prepupal
stage. Incubate the prepupae at 20°C
2) Wait the appropriate amount of time before dissecting the pupae. Pupae that areless than 40 hrs. old are more mushy and therefore more difficult, but not
impossible, to dissect than those that are older. There is an excellent discussion of
the various stages of pupal retina development described in Cagan and Ready (Dev.
Bio. 136, pgs. 346-362).
3) Remove the hard exterior cuticle by cutting the posterior 1/6-1/8th of cuticle off.
This can be done with a surgical blade by carefully cutting through only the hardexterior cuticle all the way around the pupae somewhat like using a can opener.
4) Inside the hard exterior is a varying degree of mush encased in a soft transparent
cuticle layer. After step 3 the posterior end of the pupae will be visible through thehole made in the exterior cuticle. Grab both the posterior end of the pupae and theanterior end of the hard external cuticle with #5 forceps gently pulling the pupaefrom inside the hard external cuticle. Some mush will probably leak out from theposterior of the pupae during this step but the anterior end will remain intact.
5) Place the liberated pupae immediately into PBS or 0.1 M phosphate pH 7.2 forfurther dissection. Using a surgical blade cut a hole, from dorsal to ventral, in theanterior edge of only the transparent soft cuticle. Be careful not to smash theanterior end too much. Once a small hole is created in the cuticle it can be enlarged
by inserting the tip of the surgical blade in the hole and cutting in the dorsal andventral directions.
6) With #5 forceps, give one or more forceful squeezes to the pupae starting near the
middle and working toward the anterior. After the first couple of squeezes, thebrain with attached retina and alot of mush should pop out the hole created in theanterior of the pupal cuticle. Wash the mush away with a yellow pipet tip attached
to a P20. The brain and retina will not be obvious until the mush has been washedaway. At 40 hrs. of pupal development the brain and retina look sort of like twomushroom tops attached by their stalks.
7) In order to transfer the brain retina complex from one vessel to another, spear thecomplex in the middle of the brain with a very fine tungsten needle.

63

Antibody and cobalt staining of pupal eye discs

Antibody Staining of Pupal Retina/Brain Complexes:
1) The pupal retina/brain complex is dissected as above in 0.1 M PO4 or PBS and thentransfered into an eppendorf tube containing 700 µl of fixation solution. Transfer
the complex by spearing the middle of the brain with a fine tungsten needle. Somepeople prefer to transfer the complex by pipetting it into a P20 tip. Two fixation
solutions which work well on fly retinas are (a) PLP (see eye disc staining protocol)and (b) PEMP (0.1 M Pipes pH 7.0, 2.0 mM EGTA, 1.0 mM MgSO4, 4%
paraformaldehyde). Fix the retina/brain complex for 30-40 min. on ice.
2) Remove the fixation solution by aspiration and add 1 ml of ice cold PBT (1 x PBS,
0.2% BSA, and 0.1% Triton) or PSB (0.1 M PO4, 0.1% saponin, 0.2% BSA). The PBT
buffer will provide better penetration of antibodies as triton is a stronger detergent
than saponin. However, some membrane antigens are soluablized by triton so be
careful in your choice of wash solution.
3) Wash the retina/brain complexes at least 30-40 min. with a minimum of 5 more 1
ml changes of wash solution (PBT).
4) Remove the last wash and replace it with 100-200µl of your primary antibody
dilution in PBT + 5-10% serum. The appropriate primary antibody dilution
required for your antibody must be determined empirically. Incubate retina/brain
complexes in primary antibody on ice in the eppendorf tube for at least 2 hrs.
5) Remove the primary antibody and perform 3 x 10 min. washes with 1 ml of ice cold
PBT solution.
6) Replace the last wash with 100-200 µl of diluted secondary antibody in PBT + 5-10%
serum. Our lab uses a Goat anti-mouse HRP conjugated second antibody obtained
from New England Bio Labs. We use this antibody diluted 1/250-1/500. Jackson
Immuno Research sells comparable reagents conjugated with biotin, HRP, Alkaline
Phospatase, and fluorescent chromophores. The choice of secondary antibodies
should be based on the sensitivity required and the application. Incubatecomplexes in secondary antibody for 1-2 hrs.
7) Wash the complexes 3 x 10 min. with 1 ml of ice cold PBT per wash.
8)1 During the last wash prepare the HRP staining solution which should be 1 x PBS
containing 0.5mg/ml dimethylaminoazobenzene (DAB), 0.02% CoCl2.6H2O, and
0.003% H2O2. The CoCl2 and H2O2 should be added just before use of the staining
solution. Remove the final PBT wash and add the staining solution to theretina/brain complexes. Follow the staining reaction under a microscope and stop
it by adding two 1 ml washes of cold PBS. Unused DAB solution should be
disposed of in bleach.
9) Dehydrate the stained complexes by passing them through an ethanol series (30%,50%, 70%, 90%, 2 x 100%) and then mount the retinas in DPX (Fluka Chemicals).
1 This protocol describes a staining procedure which uses cobalt intensification of the

DAB precipitate. Other intensification methods are also possible and are described in
the eye disc staining protocol.

64

Antibody and cobalt staining of pupal eye discs

Once placed in DPX the retinas can be dissected from the brain hemispheres andmounted flat.
Cobalt Sulfide Staining of Pupal Retinas:
1) Dissect the pupal retinas as described above in 0.1 M PO4. The dissection and all
the other fixing and staining steps are carried out in puddles on a Sylgard
dissecting dish. Transfer the retina/brain complexes from one puddle to the next
by spearing the brain with a tungsten needle.
2) Fix the retina/brain complexes for 5-15 min. at r.t. in 2% gluteraldehyde, 0.1 M PO4(pH 7.2).
3) Dip the speared retina in a puddle of 0.1 M PO4 briefly and then soak the retinas in
2-4% Co(NO3)2.6H2O for 5 min. at r.t.
4) Dip the speared retina/brain complex into water for a brief wash.
5) Transfer the retinas into 1-2% (NH4)2S and incubate at r.t. until the retinas turnblack. If the retinas do not turn black use the higher concentrations of the cobalt
and (NH4)2S reagents.
6) Wash the retinas briefly by dipping the speared complexes into water.
7) Mount the retinas in 80% glycerol or Aqua-Poly/Mount (Polysciences). Dissect theretina from the brain and mount it flat.
8) See Cagan & Ready (Dev. Bio. 136, pgs. 346-362) for an excellent description of
pupal retina development.
Sylgard 184 Silicone Elastomer Kit (Dow Corning) is available from
K.R. Anderson,
2800 Bowers Ave.
Santa Clara, CA 95051
408-727-2800.
You can add India ink for a dark work surface.

65

revised & edited by Marek Mlodzik

ß-GALACTOSIDASE DETECTION IN EMBRYOS
The following collection contains several protocols provided by Yash Hiromi, Christian
Klämbt and others.
ß-Galactosidase Activity Staining using X-gal
The protocol from Y. Hiromi contains several staining solutions that allow adjustment tothe staining intensity in particular genotypes. The use of cacodylate as buffer in the
fixative and DMF as solvent for X-gal can be omitted and replaced with less toxic
solutions; PBS as buffer during fixation and DMSO as solvent for X-gal.
However, the more recent protocol by Ch. Klämbt appears much easier to use (no/or
chemical devitellinization of embryos) and is certainly the one of choice when screening
large numbers of different genotypes. It also allows counter staining with antibodies
following the ß-gal activity reaction.
Protocol from Yash Hiromi
1)

Dechorionate embryos with 50% bleach. Wash well with water and collect on a
Nitex mesh. Blot mesh on tissue paper to remove excess water.

2)

In a deep depression slide, fix embryos in 0.5 ml of heptane which is saturated withthe fixative for 15 minutes at room temperature. Cover the wells with a glass slide
to prevent heptane from evaporating. Embryos should turn light yellow. Whitish
embryos are not well fixed and will not stain well.

3)

Take embryos with a Pasteur pipette and transfer onto a slide glass. Use a piece offilter paper to remove excess heptane. After all the heptane has evaporated, takeembryos by gently touching with a double stick tape and stick the tape on a slide
glass, embryo side up (Siliconized slides are easier to work with.) Cover embryoswith a drop of PBS.

4)

Devitellinize using a dissection needle (a tungsten needle is not necessary). You can
use a sewing needle whose tip is sharpened with a sand paper. Well fixed embryos
should be stiff and should come out from the membrane by gently scratching thevitelline membrane. Transfer the embryos into an Eppendorf tube.

5)

Remove the PBS and add ~300 ml of staining solution without X-gal (i.e. Fe/NaP or
Fe/CP). Leave at room temperature for ca. 5 minutes.

6)

Remove liquid and replace with staining solution. Incubate from 2 hours toovernight at room temperature or 37° C.

7)Remove the staining solution. Rinse embryos by vortexing once in 70% ethanol andonce in 100% ethanol.

8)Store the embryos in 90% glycerol/PBS at 4° C. They can be kept for months at thisstage. Mount using two #1 coverslips as spacer.

Comments:

66

ß-Galactosidase detection in embryosa)Staining with Fe/NaP pH 7.2 and higher temperature (37° C) results in stronger
staining, presumably because the conditions are closer to the optimum for the
enzymatic activity. For ftz/lacZ fusion gene transformants, staining using Fe/CP
overnight at room temperature is sufficient for visualizing stripes in the extended
germband stage embryos and for CNS staining. To stain stripes at the blastoderm
stage, use Fe/NaP pH 7.2 and stain overnight at 37° C. In this case, do not include
older embryos in the same tube, because they will stain too strongly and may
transfer the blue reaction product to the other embryos in contact. For best
resolution, try to use conditions that allows you to stain overnight, rather thanstopping the reaction after a short incubation period. Staining at 37° C will increase
the size of reaction product crystals and will make identification of stained cells
more difficult.

b)White precipitate (X-gal crystals ?) will form in the staining solution during
incubation. This does not affect staining reaction, but will stick to the embryos.
Vortexing in ethanol helps to remove it from the embryos.

c)Embryos can also be dehydrated and mounted in Epon for permanent preparations.
GMM can not be used because blue dye appears to dissolve (although slowly) inthis medium.
Solutions:Fixative (prepare fresh solution every day):
0.1 M Na cacodylate buffer pH 7.3
50% glutaraldehyde(EM grade Fluca #49631)
heptane

1 ml
1 ml
2 ml

Shake well and let the phases separate. Use the heptane (upper) phase forfixation. Fixative (lower) phase can be reused by adding heptane to 2 ml.

67

ß-Galactosidase detection in embryos
Fe/NaP-solution (pH 7.2):
0.2 M Na2HPO4
0.2 M NaH2PO45 M NaCl
1 M MgCl2
50 mM K3(Fe(CN)6)
50 mM K4(Fe(CN)6)
H2O
Store in dark at room temperature.

1.8 ml
0.7 ml
1.5 ml50 ml
3.05 ml
3.05 ml
to 50 ml

Fe/CP-solution (pH 8):
50 mM K3(Fe(CN)6)
50 mM K4(Fe(CN)6)
CP pH 8Store in dark at room temperature.

5 ml
5 ml
40 ml

CP-solution (pH 8):
0.1 M citric acid
Na2HPO4
H2O

27.5 ml27.61 gto 1 lStaining solution:
Warm up Fe/NaP or Fe/CP to 37° C. Add 1/30 volumeof X-gal (5-bromo-4-chloro-3-indolyl-b-D-galactopyranoside)solution (8% in DMSO, store at -20° C) . Use 200 to 300 ml per
tube of embryos.
Modification for weak ß-Galactosidase Expression
With weak ß-galactosidase expression, long incubation of glutaraldehyde fixed embryosin staining solution results in an undesirably yellow embryo. It is possible to avoid thiswith the following modification
1) Fix dechorionated embryos for 20 minutes in a monolayer in heptane previously
equilibrated with 18% formaldehyde in 0.1 M cacodylate buffer, pH 7.4.
2) As in other X-gal protocols, dry embryos and mount on double sided tape, inside asilicone sealant well on a slide.
3) Wash with staining buffer, then incubate for up to 3 days at 30° C in staining
solution with Xgal. Refresh solutions half way through.
4) Wash out buffer and replace with fixative (for example 4% formaldehyde in PBS).
Poke a hole in the vitelline, let fixative penetrate for at least 10 minutes, and thendevitellinate embryos. Store in 80% glycerol/20% PBS until photographed.

68ß-Galactosidase detection in embryos
Protocol from Christian Klämbt.
All steps are performed in custom made multi well plates unless otherwise noted.
I. Preparation of embryos still in their vitelline membrane:
1) Collect embryos and wash with PBT.
2) Dechorionate in 50% bleach for 4 minutes and wash for 2 to 5 minutes with PBT.
3) Blot the embryos dry and place them in n-heptane saturated with 2.5%
glutaraldehyde/PBS. The PBS/glutaraldehyde solution can be kept and used for thesaturation of fresh n-heptane for a long time (up to 8 months at 4° C).
4) Fix for 7 minutes while gentle agitating at RT.
5) Blot away excess heptane and wash embryos with PBT until they sink to the bottom
and do not stick to each other any more.
6) Wash and permeabilize for another 2 hours with several changes of PBT (Even slight
contamination with heptane on the vitelline membrane will cause diffuse blue
background staining.)
II. Staining:
1) Heat the staining solution to 65° C, or over a flame until the solution gets cloudy (Thiswill greatly reduce the formation of crystals during the staining process). Add 1/50
volume of 10% X-gal solution in DMSO to the warm staining solution.
2) Blot away excess PBT and place embryos in the staining solution.
3) Stain for 2 to 4 hours at 37° C in a humid chamber and stop the reaction by washingwith PBT.
III. Preparation of devitellinized embryos for X-gal staining:
Methanol and also ethanol will destroy lac-Z activity. However, if the devitellinization
step is done fast, the methanol as well as the ethanol do not significantly destroy the
enzyme's activity.
1) Wash, dechorionate and fix the embryos as in (I.).
2) Blot away excess heptane fix and wash the embroys in n-heptane.
3) Transfer the embryos in 1 to 2 ml heptane into a 5 ml glass tube, add 1 to 2 ml
methanol.and shake vigorously.
4.)After 10 to 20 seconds the interphase becomes visible. Aspirate the heptane phaseand half of the methanol phase off (at this point not all the embryos will have madeit to the bottom of the glass tube. You will lose 30 to 50% of the embryos).
5) Add 2 ml methanol for a wash. Aspirate all methanol and wash twice with 80%
ethanol. Do not let the embryos sit in methanol or ethanol for longer times thanneccessary.

69

ß-Galactosidase detection in embryos

6) Wash 3 times for 5 to 10 minutes with 4 ml PBT on a rocking platform.and wash foran additional 2 hours in PBT.
7) Transfer the embryos back into a multi well plate and proceed with the staining as in(II.). Do not let the embryos dry at this point. If you want to look at the X-gal stained
embryos, wash briefly in PBT and mount them in PBS/50% glycerol.
If counter staining of the embryos with specific antibodies is desired, wash awaystaining solution with several changes of PBT for 2 hours. Subsequently wash embryoswith PBT containing BSA and 5-10% goatserum for 1 h and proceed with the usual
staining protocol for antibodies.
Solutions:
Staining solution (200 ml):
10 mM PO4-buffer pH 7.2
150 mM NaCl 6.0 ml 5M
1 mM MgCl2
3 mM K4[FeII(CN)6]
3 mM K3[FeIII(CN)6]
0.3 % Triton X-100
PBT:

2.0 ml 1M
0.2 ml 1M
0.3 g
0.2 g
0.6 ml

PBS with 0.5 % Triton X-100 (but without BSA)

Visualization of ß-Galactosidase for EM Histology (Protocol from Roger Jacobs)
1) Process for X-gal staining as described above for light microscopy. After staining,transfer to 0.1 M cacodylate buffer, pH 7.4, and select the embryos you wish toembed.
2) Fix in primary EM fix (2% glutaraldehyde, 2% paraformaladehyde in 0.1 M
cacodylate buffer, pH 7.4) for 30 minutes.
5) Wash in cacodylate buffer with at least 3 changes for 5 minutes.
6) Fix in 1% osmium tetroxide in cacodylate buffer for 30 minutes7) Wash in cacodylate buffer, followed by distilled water.
8) Stain in 5 % uranyl acetate for 30 minutes.
9) Dehydrate and infiltrate as in standard EM protocol. Older whole-mount embryos
need longer infiltration times. Infiltrate in pure plastic for 24 to 48 hours (Notedissections infiltrate through a graded ethanol-plastic series!). Use for example
Epon-Araldite plastic. You may also use methacrylate based plastics, but this will
make semi-thin sectioning (0.5 µm) more difficult. It is much easier to embed wholemount embryos in the middle rather than the bottom of the block. Cook plastic inhalf filled molds for about 4 hours, then add embryos in new plastic to the top half ofthe mold. You can orient the embryos in the mold with a fine probe.
Sectioning and Staining:70

ß-Galactosidase detection in embryos

Trim and orient your tissue by conventional technique. Sections of 1.0 µm or thicker canbe picked up individually off a dry glass knife and placed on a small drop of water on asubbed (1% gelatin and 0.1% chromium potassium sulphate x 12 H2O) slide. For serial
sectioning, use a glass knife with a small water boat behind it [made with mylar tape, or
LKB's prefab plastic boats (#2208-100)]. 0.5µm is the easiest to handle. Ribbons ofsections can be spread with a chloroform wick, and transferred to a dry subbed slideusing the wire loop technique. Merely touch the loop containing the ribbon and water
to the slide, and remove the water by touching a sliver of filter paper to the edge of theloop.
Best tissue contrast is obtained with 0.5% Toluidine Blue O in 1% Borax buffer.
This stain is preferred when you are trimming and orienting, but will not allow you tovisualize X-gal product.
To preserve contrast with X-gal, counterstain with saturated aqueous Basic
Fuchsin. This stain penetrates epon slowly, but there are two tricks which will increase
permeability. First, you might soak your unstained slide in xylene for 20 to 30 minutes,
wash in acetone, and dry before staining. Secondly, add a drop or more of acetone
containing 1% Basic Fuchsin to the slide when staining with saturated aqueous Basic
Fuchsin on a hotplate (65° C).
Slides should be permanently mounted with permount and coverslip, an viewed
under oil immersion optics. You can now enhance X-gal staining contrast by using
appropriate filters (for example a yellow filter or in extreme cases, a red filter will
darken the image of the crystals relative to the rest of the tissue).
In EM, X-gal crystals make large ugly needles that actually penetrate cell walls,
and fill vacuoles inside cells. It is therefore useful mostly in low magnification work.

71

ß-Galactosidase detection in embryos
Immunohistochemical Detection of ß-Galactosidase
The detection of ß-galactosidase with an antibody and immunohistochemical staining
gives the best possible single cell resolution. However, it is more laborious than activity
staining.
I usually follow the standard protocol for immunohistochemistry in embryos as
described in the relevant section of this methods book. The following reagents anddilutions are normally employed:
primary antibody:anti-ß-gal monoclonal from Promega
use at 1/200 to 1/500 dilution in PBT(PBS, BSA,
Triton; for recipe see immunohistochemistry protocol)cont. 5% goat serum(The quality of the anti-ß-gal monoclonal is obviously
important. However, we have sometimes had problems
obtaining consistent results due to lower quality of the
primary antibody. Nevertheless, Promega still provides
the best monoclonal commercially available.)secondary antibody

HRP-conjugated goat-anti-mouse polyclonal from Biorad
use at 1/200 dilution in PBT/goat serum (see above)

HRP reaction

0.5 mg/ml DAB in PBT
add 1µl of 3% Hydrogenperoxide and 8µl 8% NiCl(2)7H(2)O for each ml of DAB solution

Embryo pretreatment, incubations, washes, staining reaction and mounting of theembryos is performed as described by the standard immunohistochemistry protocol.

72

Marek Mlodzik

ß-GALACTOSIDASE DETECTION IN IMAGINAL DISCS
The following protocols for ß-Galactosidase activity staining with X-gal andimmunohistochemical staining in 3rd instar larval imaginal discs are based on the
protocols for detection in embryos.
Activity staining using X-gal
1)

Wash 3rd instar larvae (wandering stage) in PBS for a few minutes. Dissect in PBS.

2)

Dissect anterior mouth complex by grabbing larvae at each end (Use Dumont #5
forceps). Hold larva at the posterior end and pull at the anterior end holding themouth parts with the forceps. Usually, tissue still attached to the mouth hooks
contains (most) imginal discs (the wing discs get sometimes lost), the brain/CNS
complex, the salivary glands, some fatbody and anterior parts of the gut. Dissect
about 5 to 6 larvae for each genotype.

3)

Leave all this tissue with the mouth hooks (it can be dissected at the end of thestaining procedure) and fix tissue in 1% glutaraldehyde in PBS for 15 min at RT.

4)

Remove fix and wash tissue with PBS for 10 min, 2 changes of buffer.

5)

Prewarm staining solution without X-gal for 10 min at 37 C.

6)

Add X-gal (1/30 of a 8% stock in DMSO) to staining solution and incubate tissue inthis solution for several hours to overnight, use 100-200µl per reaction (genotype).

7)

After staining is completed remove staining solution and rinse discs once with PBS(optional) and resuspend them in 80% glycerol/PBS. Leave tissue for a few hours inglycerol solution before mounting.

8)

Transfer tissue in a drop of glycerol (with cut off yellow tip) onto slide. Dissect
away unwanted tissue (e.g. mouth hooks) before mounting under a cover slip. For
permanent storage seal cover slip with nail polish.

Comments:
Some people prefer NaPO(4)-Buffer (pH 7.4) to PBS in dissecting and wash steps.
The staining solutions are the same as for the embryos (see embryo ß-gal protocol).
Due to the small overall size of the eye disc and the small cell size in the disc, diffusionof the ß-gal staining is more a problem in discs than in embryos.
In lacZ-lines with strong expression the following alternative staining solution can beused; it reduces diffusion but also reduces the sensitivity of the staining.
Alternative staining solution (based on A. Fire's protocol, moidified by U. Gaul):
1 ml of solution contains:

695 µl H(2)O
1 µl
MgCl(2), 1M

73

Β-galactosidase detection in imaginal discs

200 µl100 µl
4 µl

NaPi (0.5m, pH 7.2)
Redox. Buffer 100mM K(3)Fe[CN]6; K(4)Fe[CN]6
1% SDS (made fresh from 10% stock)

immediately before staining add 1/30 vol of 8% X-gal
Immunohistochemical Detection
Eye imaginal disc:
Dissections are performed as described in eye disc antibody staining protocol.
The antibody incubations and staining are also virtually identical to the protocol for eye
disc antibody stainings (see specific section).
The primary antibody, anti-ß-gal monoclonal (from Promega), is used at 1/200 to 1/500
dilutions; the secondary, HRP conjugated goat-anti-mouse (from BioRad) is used at thesame dilutions. For the incubations and washes, I usually use PBT (PBS, BSA, Triton X100; see embryo protocols) instead of 0.1m NaPO(4), 0.1 % saponin. However, there isno significant difference in staining intensity or background between those twoalternatives.
There are several options for intensification of the signal during and after theperoxidase/DAB staining reaction. The eye disc antibody staining protocol describes theosmium tetroxide postfixation and the use of Co and Ni in the DAB reaction (for details
see eye disc staining protocol). I prefer to add just NiCl(2) to the DAB reaction (noCobalt) as described in the ß-gal detection protocol for embryos. Since this enhancement
method does not postfix the tissue, an ethanol dehydration serie, 30%, 50%, 70%, 90%
and 100% EtOH, each for 10 min, is necessary to avoid shrinking of the tissue (this isquite important because the disc is already small enough!). A postfixation inglutaraldehyde as an alternative is only suitable for strongly stained discs, because partof the signal fades out during this process.
The dehydrated tissue can be stored in 100% ethanol for several days. Mounting of thediscs in DPX mounting medium is as described in the general eye disc staining protocol.

74

M. Fortini

ADULT HEAD FROZEN TISSUE SECTIONSß-GALACTOSIDASE ACTIVITY STAINING
1)

Embed up to 12 fly heads in O.C.T. Tissue Tek compound (Miles Scientific) under adissecting microscope. For horizontal sections, which are generally the best for
examining compound eye and optic lobe structures, the heads should be oriented
upright in the block, so that the ocelli are pointing towards the microscope
objectives. Slow freezing by periodic immersion of the block support in a dry
ice/ethanol bath allows several heads to be precisely positioned in a single blockwith fine forceps.

2)

Pretrim the block and place it in the cryostat for 20-30 minutes prior to cutting toequilibrate the block to the cryostat temperature. Cut 10-14 µm sections on thecryostat and transfer section ribbons onto slides (subbed slides not necessary). All
the sections from one block may usually be fit on a single slide.

3)

Warm the slides at 50-60°C for 2-3 minutes and then allow them to air dry for about
1 hour. In my experience, slides may be kept for several days sitting on a benchtop
at this stage with no apparent loss of enzyme activity.

4)

Fix the tissue sections in 2.0 % glutaraldehyde in PBS for 15-20 minutes at room
temperature in a coplin jar.

5)

Wash the slides two or three times in PBS for 5 minutes/wash.

6)

After the last wash, remove excess PBS with a tissue and place the slides on a levelsurface in a moist airtight chamber. A Tupperware box with water-soaked blotting
paper works well for this purpose. To keep the slides off the paper, they should beplaced on two lengths of plastic 5 ml or 10 ml pipettes which rest on top of thepaper block.

7)

Allow the remaining PBS on the slides to dry (5-10 min at room temp). The slides
may be left to dry for a few hours at this point with no apparent loss of enzyme
activity.

8)

Apply 50 µl of a prewarmed staining solution + X-gal to each slide and cover with acover slip (50 µl suffices for a 22 x 40 mm cover slip - adjust accordingly for different
sizes).

75

Adult Head Frozen Tissue Sections

Staining solution:
50 ml total(store at room temperature in a foil-wrapped 50 ml
conical tube in the dark - stable for months).
1.8 ml of 0.2 M Na2HPO4
0.7 ml of 0.2 M NaH2PO4
1.5 ml of 5 M NaCl
50 µl of 1 M MgCl2
3 ml of 50 mM K3(Fe(CN)6)
3 ml of 50 mM K4(Fe(CN)6)
bring to 50 ml with ddH2O
X-gal 30X stock solution: store at -20°C
10% X-gal in N,N-dimethylformamide
IMPORTANT: To avoid instantaneous precipitation of X-gal crystals when applying
solution to slides, it is necessary to prewarm these solutions. Generally, while theslides are in the fixative or washes, I aliquot out the required amount of staining
solution into an eppendorf tube and incubate in a 42°C temp block for about 5
minutes. I then add 1/30 volume of the X-gal stock solution to this tube, quickly
vortex or shake, and place it back in the 42°C temp block. I apply the staining
solution + X-gal onto the dried slides as quickly as possible, put on all the coverslips, and immediately put the sealed chamber in a 37°C incubator.
9)

Allow the staining reaction proceed for as long as necessary (from a few minutes toovernight).

10) Examine the slides periodically under a dissecting microscope. When the staining
appears to be complete, float the cover slips off in PBS and wash the slides 2 x 5
minutes in PBS.
11) At this point, the slides may be dehydrated through a graded ethanol series andmounted in DPX (Fluka) or other suitable mountant. The ethanol series also helps
remove X-gal crystals deposited on the tissue sections during the staining reaction.
Alternatively, the slides may be simply mounted in 70% glycerol in PBS after the
final PBS wash (in my experience, the glycerol mounting gives better morphology
for photomicroscopy but is less suitable for long-term storage of slides).
12) For Hoechst labeling of nuclei, do not dehydrate the sections. Instead, pipette 0.51.0 ml of a 1 µg/ml solution of Hoechst 33258 in PBS directly onto each slide. After
1-2 minutes, drain the dye solution off, rewash the slides in PBS a few times (5minutes each) and mount in 70% glycerol/PBS.

76

Adult Head Frozen Tissue Sections

ANTIBODY STAINING
The protocol provided below has been found to work well for antibody stainings offrozen sections of Drosophila head tissues, and it should also be suitable for staining ofother tissues. The sucrose infiltration step (2) provides improved cryoprotection of thetissue sections but may be omitted if good morphology of the tissue is not required.
1)

Fix the tissue of interest in 2.0% formaldehyde in PBS at 4°C. For Drosophila heads,
first remove the proboscis and fix the head capsule for 60-90 minutes. For othertypes of tissues, adjust the fixation time according to their relative size andestimated efficiency of infiltration.

2)

Wash the tissue in PBS and transfer to 12% sucrose in PBS at 4°C. Allow thesucrose solution to infiltrate the tissue for 16 hrs.

3)

Remove the tissue from the sucrose solution and submerge it in a drop of O.C.T.
Tissue Tek Compound (Miles Scientific). Allow the tissue to be permeated by the
O.C.T. compound for 10-30 minutes at room temperature and then embed thetissue in frozen O.C.T. compound using an ethanol-dry ice bath (see preceding
protocol.)4)

Place the frozen block containing the tissue to be sectioned in the cryostat chamber
for at least 20 minutes to equilibrate it to the cryostat temperature (-14°C to -18°C).

5)

Cut 10-14 µm sections on the cryostat and load onto freshly gelatinized slides.

6)

Heat slides briefly at ~40°C on a drying plate for no more than one minute to dry
the sections onto the slides. Overheating may adversely affect the antigenicity ofthe protein of interest and the morphology of the tissue.

7)

Fix the sections immediately in 0.5% formaldehyde in PBS for 20-60 minutes atroom temperature. Slides may be stored in this fixative at 4°C for a few days ifnecessary.

8)

Wash the slides two or three times for 2-3 minutes/wash in PBS.

9)Block the slides for 30 minutes in PBSG (0.2% BSA, 1% goat serum, 0.01% saponin
in PBS).

10)

Wash the slides through several changes of PBS/0.01% saponin.

11)

Apply 75-150 µl of the primary antibody at the appropriate dilution in PBSG toeach slide. Unless an excess of antibody solution is used, gently place cover slips
on the slides to evenly spread the antibody solution over the tissue sections (100 µlof solution suffices for a 22 x 40 mm cover slip). Incubate the slides in the primary
antibody for 30-60 min at room temperature or overnight at 4°C in a moist, airtightchamber (ie. a Tupperware box containing wet blotting paper).

12)

Wash the slides through several changes of PBS/0.01% saponin (float off the coverslips in the first wash).

77

Adult Head Frozen Tissue Sections

13)

Incubate the sections with the secondary antibody at the appropriate dilution in
PBSG exactly as described in step 11 for the primary antibody. For Bio-Rad
Laboratories HRP-conjugated goat-anti-mouse IgG, a 1:200 dilution for 30-60
minutes at room temperature works well.

14)

For an HRP-conjugated secondary antibody, the slides are again washed throughseveral changes of PBS/0.01% saponin and then incubated in 0.5-1.0 ml of staining
solution per slide without cover slips or 100-150 µl per slide with cover slips. The
staining solution consists of: 0.5 mg/ml diaminobenzidine (DAB), 0.003% H2O2,
1.5 mM CoCl2, and 1.5 mM NiCl2. The CoCl2 and NiCl2 are optional components;
in their absence, a brown product forms, whereas in their presence, a more intense
blue-black product is obtained. Since the extent and localization of backgroundstaining can vary considerably between intensified and non-intensified
preparations, it is often worth trying both methods in parallel.

NOTE:

DAB is thought to be a potent carcinogen and is to be handled with care. All DABcontaining tubes, pipette tips, used DAB solutions and similar materials should be
immersed overnight in 100% bleach and disposed of according to the proper
regulations.

15)

Monitor the staining reaction under a dissecting microscope. When the reaction
has proceeded to completion (usually 5-30 minutes), it is stopped by washing theslides through a few changes of PBS.

16)

For compound microscope viewing and photomicroscopy, the sections may either
be dehydrated through a graded ethanol series and mounted under cover slips in astable mountant such as DPX (Fluka Biochemie), or they may be mounted undercover slips in 70% glycerol/PBS after air-drying without dehydration. In general,the glycerol mounting is superior for photomicroscopy but is not suitable forstable, long-term storage of samples.

17)

For Hoechst flourescent counterstaining of nuclei, do not dehydrate the sections
after step 15. Instead, pipette 0.5-1.0 ml of a 1.0 µg/ml solution of Hoechst 33258 in
PBS directly onto each slide, allow to sit for 1-2 minutes, then rinse off through a
few changes of PBS and mount in 70% glycerol/PBS as described above.

78

Yash Hiromi

WHOLE MOUNT STAINING OF DROSOPHILA RETINA FOR
β-GALACTOSIDASE ACTIVITY
1)

Prepare postfix solution in wells of a microtiter plate, and fixative-saturated
heptane in a well of a deep depression slide.

2)

Fix whole adults in 0.5ml of fixative-saturated heptane for 20 minutes at room
temperature. Cover the wells with a sideglass to prevent heptane from evaporating.

3)

Remove adults from heptane and place on a piece of blotting paper for a few
seconds to evaporate heptane. Place in a drop of postfix solution in a shallow
depression slide. Remove body from the head and discard.

4)

Using two tungsten needles, dissect retina from head as follows. First, cut along theedge of the lens and remove retina + cornea from the rest of the head. In most casesa thin pigmented layer of cells located between the retina and the lamina will come
off the retina. Since in many lacZ lines this layer stains heavily and masks theexpression in the retina, it needs to be removed by scratching with a fine needle.
Then, remove the lens from the retina by inserting the needle between the retina
and the cornea. Removal of the cornea is not absolutely necessary for staining, but
it accelerates bleaching of the pigments and will also make the preparationhydrophilic. Transfer the dissected retina to postfix solution in a microtiter plateand fix for an additional 10 minutes.

5)

Remove the postfix solution using a drawn out Pasteur pipet and rinse once with
PBS, and then replace the PBS with staining solution without X-gal (i.e., Fe/NaP or
Fe/CP). Leave at room temperature about 5 minutes.

6)

Remove liquid and replace with staining solution. Incubate 2 hours to overnight atroom temperature or 37°C.

7)

Remove stianing solution. Rinse retina once in 70% ethanol and once in 100%
ethanol. This step helps to remove X-gal crystals that are adhering to the sample.

8)

Remove ethanol and add 50% glycerol/PBS. Take retina out into a drop of 50%
glycerol in a depression slide and remove X-gal crystals with a fine needle. Mountin 50% glycerol using one #1 coverslip on one slide.

Staining with Fe/NaP pH7.2 and higher temperature (37°C) results in stronger staining,presumably because the conditions are closer to the optimum for the enzymatic activity.
For Rh4/lacZ fusion gene transformants, using Fe/NaP overnight at room temperatureallows you to stain overnight, rather than stopping the reaction after a short incubation
period. Staining at 37°C will increase the size of reaction product crystals and will makeidentification of stained cells more difficult.
Fixative-saturated Heptane (Prepare fresh solution every day)
0.1M Na cacodylate buffer pH7.3
1ml
50% gluteraldehyde
1ml(EM grade, Fluka #49631)
heptane2ml

79

Whole-mount staining of drosophila retina for β-galactosidase activity

Shake well and let the phases separate. Use the heptane (upper) phase forfixation. Fixative (lower) phase can be reused by adding heptane to 2ml.
Postfix Solution
0.1M cacodylate buffer pH7.32ml
water
50% gluteraldehyde(EM grade, Fluka #49631)
10% Triton X-100

2ml
80µl (1% final concentration)

Fe/NaP (pH7.2)
0.2M Na2HPO4
0.2M NaH2PO4
5M NaCl
1M MgCl2
50mM K3(Fe(CN)6)50mM K4(Fe(CN)6)
H2O

1.8ml
0.7ml
1.5ml
50 µl
3.05ml3.05mlto 50ml

20µl (0.05% final concentration)

Store in the dark at room temperature.
Fe/CP (pH8)50mM K3(Fe(CN)6)50mM K4(Fe(CN)6)
CP pH8

5ml
5ml
40ml

Store in dark at room temperature.
CP (pH8)
0.1M citric acid
Na2HPO4
H2O

27.5ml27.61gto 1 liter

Staining Solution
Warm up Fe/NaP or Fe/CP to 37°C. Add 1/30 volume of X-gal (5-bromo-4chloro-3-indolyl-β-D-galactopyranoside) solution (8% in DMF (N,Ndimethylformamide), store at -20°C).

80

Kevin Moses

EMS MUTAGENESIS
EMS(ethane methyl sulfonate or methanesulfonic acid ethyl ester as Sigma calls it) A
chemical mutagen that most commonly causes transitions by methylation of G residues
but can yield a spectrum of mutations such as deletions etc. Mr = 124.2g ρ = 1.17 g/ml.
EMS can be ordered from Sigma (catalogue No. M-0880).
SafetyEMS is a mutagen and a carcinogen. It should therefore be handled in a hoodwith gloves. EMS solutions can be deactivated in a solution of 4g NaOH and 0.5 ml
thioglycolic acid in 100 ml. We store contaminated dry waste in sealed plastic bags
under a hood, which are periodically removed by University safety officers.
Screen design
Normally one treats males and then crosses them to untreated female virgins. Ina simple screen these females will be marked with the mutation of interest and the F1progeny can be screened for new alleles. There are many more sophisticated types ofscreen. In order to distinguish the chromosome bearing the new allele from the maternal
chromosome one of these may be marked with another (at best closely linked) mutation,ideally a recessive also present on the appropriate balancer. It may be desirable to screen
only the F1 sons in order to avoid recombination between the marker and new allele. It
is best to screen for some other RK1 marker while screening for the gene of interest,(probably the best are eye colour mutations such as white, cinnabar or scarlet) to give you
something to find and as a running positive control for the effectiveness of thetreatment.
Males
It is best to use more males than you would normally as their virility can beaffected by the treatment. Avoid the marker yellow, which reduces virility and increases
mortality during treatment. Ideally treat 3 day old males. Those that are too young will
not yet be fertile, those that are too old may suffer from Portnoy's complaint.
DosageDosage is controlled by EMS concentration and can be monitored by scoring for
X-linked lethals. This is best done by crossing some of the treated males en masse tovirgins bearing an X-chromosome balancer (such as FM6). The resulting F1 daughters
are allowed to mate freely to their brothers, and after several days are placed in tubes
individually and allowed to lay for a few days, and are then discarded. The F2 sonsmust inherit the X-chromosome from their F1 mothers which are now a single isolates(called *) from the original mutagenised grand-fathers. The F2 males can thus be either
FM6/Y or */Y. In some lines * can carry a newly induced lethal mutation and thus only
FM6/Y sons will appear. Thus the % of lines with no Bar+ F2 sons is the % newly
induced X-linked lethals. Typically 25 mM EMS will yield about 30% X-linked lethals,
which equates to roughly one hit per autosome on average, and 1 hit in 2000 to 5000 for
most loci. The effective dosage of EMS may vary with different lab conditions or stocks,
so a pilot run to test several EMS concentrations may be worthwhile.
Mosaicism
Orthodoxy holds that EMS mutated post-meiotic sperm undergo DNA repair inthe embryo. This may well occur after the first cleavage division so it is quite common

81

EMS Mutagenesis

for the F1 progeny to be somatic and sometimes germ-line mosaics for a new allele. Thus
often apparently mutant F1 individuals can yield a mixture of F2 progeny, or no F2
individuals bearing the new allele. This is tiresome but unavoidable, and the rate of such"false positives" will depend in part on the distance on the zygotic fate map between theorgan which is the focus of your mutation (e.g. the eye for white mutations) and the germline. This rate of "false positives" can easily be 50%.
Clusters
If progeny are recovered from cells that were pre-meiotic during EMS treatment,
several identically mutant sperm could result from their subsequent mitotic division.
Thus the same allele could be recovered multiply. As spermatogenesis requires 5 days at
25°C, so long as the parents are discarded on the fourth day after treatment this should
not occur. It may be that the mutagenesis rate is not constant in these different daily
cohorts of sperm.
References
These topics are well discussed in Tom Grigliatti's chapter 2 Mutagenesis in David
Roberts' book Drosophila a practical approach (IRL press, Oxford, Washington DC 1986),
and in Michael Ashburner's Drosophila, A Laboratory Handbook (Cold Spring Harbor
Press, Cold Spring Harbor New York, 1989).
Treatment :1) Make solution I (100mM Tris Ph7.5 and 10% sucrose). Autoclave and store at room
temperature.
2)

On the day of use make deactivating solution (4g NaOH + 0.5 ml thioglycolic acidin 100 ml). Put this in the hood along with plenty of paper towels etc. and a plastic
bag for dry waste such as gloves and yellow tips.

3)

In the hood, on a diaper, dilute solution I 1+9 and add EMS. 26.5 µl EMS in 10 mlyields a 25mM EMS solution.

4)

Make up glass tubes with wadded kimwipes or filter papers in the bottom, andcotton bungs on top. Make two such for every 100 males to be treated.

5)

Transfer the males to empty bunged vials and allow to dehydrate for 30 minutes.

6)

In the hood, on a diaper, pour the EMS solution from tube to tube so as tothoroughly soak the paper in half of your prepared tubes and dispose of the residue
by dilution in deactivating solution. Do not allow the cotton bung to become wet(this will prevent the tube from ventilating when the males are in it and they will
die).

7)

In the hood, on a diaper, transfer the dehydrated males to the EMS tubes, and leavethem there over night. Leave a sign up on the hood to notify others of the presenceof EMS.

8)

In the morning transfer the males on to the dry tubes to allow them to wipe theirfeet and hopefully shit-out most of the EMS.

9)

After 30 minutes transfer the males to freshly yeasted ordinary fly food tubes andproperly dispose of all of the dry waste. The males may now be removed from thehood. Allow them a couple of hours to recover, then cross them.

82

EMS Mutagenesis10) Sub on the crosses every day for four days and discard the adults on the fifth.

83Marek Mlodzik

LOCAL MUTAGENESIS BY IMPRECISE P-ELEMENT EXCISION
This method is based on the fact that in most cases P-elements excise imprecisely
when transposing via the P transposase. Most of these imprecise excision events leave
sequences from the P elements behind. However, in a reasonable proportion of excisions(~10%) flanking genomic DNA is removed with the P elements, creating small deletions
around the original P element insertion point.
The use of this approach allows to create local mutations in genes neighboring
the original P insertion, e.g. in lines of the P[lacZ] insertions that do not have aphenotype by itself.
The following crossing schemes are using the ∆2-3 element at 99B as a source ofthe transposase. The given schemes are designed for P-elements with rosy as selectable
marker. Nevertheless, very similar schemes can be used for other common markers suchas the white gene.
II. Chromosome P Insertions
1st generation:MMP[ry+] / P[ry+] ; ry / rymmCyO / Sp ; Sb, ∆2-3 / TM6, Ubx

x

2nd generation (here occurs the excision):
M (single males)P[ry+] / CyO ; Sb, ∆2-3 / ryxmm
CyO / Sco ; ry / ryxmm
CyO / Sco ; ry /ry

3rd generation:
M (single males, ry eye color)
E{P} / CyO; ry / ry

4th generation (brother-sister cross):
MM
E{P} / CyO; ry / rymmE{P} / CyO ; ry / ryxestablished stock, if excision is lethal; check homozygous flies for visible phenotypes.

84Local mutagenesis by imprecise P-element excision

III. Chromosome P Insertions
1st generation:MMP[ry+] , ry / P[ry+] , ryxmmKi, pp, ∆2-3 / Ki, pp, ∆2-3xmm
TM3, ry / rf10 (ftz, e, ry)xmmTM3, ry / rf10xmmE{P}, ry / TM3, ry

2nd generation (here occurs the excision):
M (single males)P[ry+] , ry / Ki, pp, ∆2-3

3rd generation:
M (single males, ry eye color)
E{P}, ry / TM3, ry

4th generation (brother-sister cross):
MM
E{P}, ry / TM3, ry

established stock, if lethal; check for visible mutations in homozygous flies

85Local mutagenesis by imprecise P-element excision

X Chromosome P Insertions:
1st generation:mmP[ry+] / P[ry+] ; ; ry / ryMM+ / -- ; ; Ki, pp, ∆2-3/ Ki, pp, ∆2-3x

2nd generation (here occurs the excision):
M (single males)P[ry+] / -- ; ; Ki, pp, ∆2-3 / ryxmm
FM6 / dnd ; ; ry / ryxMMFM6 / -- ; ;3rd generation:m (single virgins, ry eye color)
E{P} / FM6 ; ; ry / ry

ry / ry

established balanced stock, if excision is lethal; check for visible phenotypes inhomozygous flies.

86Local mutagenesis by imprecise P-element excision

Richard Carthew

X RAY MUTAGENESIS

Collect 0 to 2 day old adult males and let them age a further 2 days at 25 C.
Aliquot the flies into groups of 50 each. Place each group of 50 into a gelatin capsule(Lilly, No.000) which has been perforated with a 23 gauge needle. Put the capsules in apetri dish on ice. Make sure the flies get cool. Take care that the flies and capsule do
not get wet. Place the dish in the X ray machine and give the flies a 4000 rad dose (115kV; 5 mA). Alternatively, place males in empty vials and lay vials on their sides in xray machine before giving them a 4000 rad dose. Immediately transfer the flies to a vialwith plenty of yeast and allow them to recover for a few hours. Let the males mate withan equal number of 3 day old virgin females. I like to put 30 of each sex in a bottle.
Knock the flies into fresh bottles on successive days and on the fourth day discard theflies. Starting with 400 males, one can easily get ten to twenty thousand F1 flies toscreen.

87

Richard Carthew

MICROSCOPIC ANALYSIS OF THE ADULT RETINA
Fixation and embedding for microscopy:
Dissect the head away from the body. With a number 11 scalpel blade, gently cut
away one of the eyes to expose the inside of the head. Avoid putting pressure on thehead when cutting (making the other eye 'bulge'). Place the dissected head into 2%
glutaraldehyde in phosphate buffer. This solution should be on ice. Often it isnecessary to tap the head down into the solution as it has a tendency to float.
Alternatively, you can give them a 5 second spin in a microfuge to submerge it.
Add an equal volume of 2% OsO4 in phosphate buffer. Only handle OsO4 in thehood and wear gloves; it is volatile and extremely toxic. Incubate the mixture for 30 min
on ice. Remove the glutaraldehyde/OsO4 mixture and wash the tissue with cold
phosphate buffer by filling the tube completely. Dispose of waste glutaraldehyde andOsO4 by adding an excess of dry milk powder to the waste solutions in a bottle. The
deactivated solution can then be safely removed. Add fresh OsO4 to the tissue andincubate on ice for 1-2 hours. Discard the OsO4 and dehydrate the tissue by successive
10 min incubations on ice in the following ethanol solutions: 30%, 50%, 70%, 90%, 100%.
For preparing tissue for electron microscopic analysis, also include an 80% EtOH wash.
Repeat the final 100% EtOH incubation.
Replace the final alcohol treatment with propylene oxide and incubate for 10 min
at room temperature. Handle propylene oxide with gloves and only use it in the hood.
Repeat the propylene oxide wash twice. Add an equal volume of durcapan resin to thelast propylene oxide wash and thoroughly mix. Handle resin with non-latex gloves as itis carcinogenic when unpolymerized. Incubate overnight at room temperature. Use soft
resin for semi-thin and thick sectioning and use hard resin for thin sectioning.
Replace the resin/propylene oxide mixture with pure resin and incubate at least
4 hours. Place tissue in a drop of resin on a slide. Examine the head under a dissecting
microscope and discard any poorly fixed heads. Dissect away unwanted tissue from thehead. Using a needle, transfer a fixed head to a mold which has been filled with resin.
Useful molds are BEEM or silicone rubber flat embedding molds (Ted Pella Co.). Placethe head very close to the edge of the mold (Figure 1). Orient the head so that the head isanterior up and the eye to be sectioned is closest to the edge. Make sure that the head isflat and resting on the bottom of the mold. Bake the resin at 70°C overnight. Bake any
waste resin before discarding.

Figure 188Microscopic analysis of the adult retina

Sectioning and Staining Specimens.
Examine the embedded heads to ensure that they did not shift position while theresin hardened. Only section those which maintained their correct orientation. Trim theresin block with a razor blade, preferably the kind which are Teflon coated. Trim away
excess resin which covers the eye. This is diagrammed in Figure 2. Start by trimming
resin to form the face of the block which will be cut. Then cut away resin to form a
trapezoid shape, so that the final form resembles a pyramid with its top lopped off.

Figure 2
Cut sections with your favorite microtome. Section thickness can range from 0.5
to 5 microns for tissue embedded in soft resin, and from 50 to 300 nm for tissue
embedded in hard resin. Sections can be collected either with a wire loop or a shaved
wooden stick. The stick is best for collecting serial sections when section order must bepreserved.
Transfer the sections to a puddle of water on a subbed slide. Place the slide on aheating block (about 80°C) and allow the water to evaporate. If you wish to examine
unstained sections, add a drop of DPX (Fluka) and a cover slip. If you want to stain theeye sections, use toluidine blue. Place slides with sections on them onto a heating block
which is at a temperature of 70-80°C. Allow the slides to warm up. Add several drops of
toluidine blue solution (enough to only cover the sections) to the slide, and incubate. The
incubation time is variable: it depends on the temperature of the block, how much
toluidine solution is added, etc. Therefore, make an empirical test by staining for several
different amounts of time. Around 30 seconds is usually optimal. After incubation,
remove the slide and rinse thoroughly with water Examine under the microscope for
how well the staining worked. If necessary, you can repeat the procedure until staining
is optimal. Take care; it is possible to overstain tissue with this procedure. Allow theslides to completely dry before mounting in DPX.

89

Microscopic analysis of the adult retina

Solutions:
1) Phosphate buffer. This is 0.1 M Sodium Phosphate pH 7.2. It can be made by mixing
0.2M Na2HPO4 and 0.2M NaH2PO4 in a 72:28 ratio, respectively, and then adding anequal volume of water.
2) Durcapan Resin. This can be ordered as Durcapan ACM from Fluka. It is based on
the formula for Araldite resin, and is stored for long periods of time at room
temperature as four separate components: epoxy resin (A), hardener (B), accelerator(C), and plasticiser (D). These components can be mixed together and either used
immediately or stored in aliquots at -20°C. If stored at -20°C, you can freeze-thaw analiquot for up to three times without affecting the resin. The resin is quite toxic, sobake any waste at 70°C overnight before discarding. The recipes for soft and hardresin are:
SOFT
(grams)HARD(grams)

54
44.5
2.5
10

50
50
1.75
0.75

Resin A
Hardener B
Accelerator C
Plasticiser D

3) Glutaraldehyde and Osmium Tetroxide can be purchased as concentrated solutions
from Ted Pella Co. and diluted into phosphate buffer. Store both solutions at 4°C in asealed container. 2% glutaraldehyde solutions are good for one month, and 2% OsO4
solutions are good for several months. See page 73 for instructions on disposal ofwaste glutaraldehyde and OsO4.
4) Toluidine Blue Solution. Dissolve toluidine blue and borax to 1% final concentrationfor each in water. Store it in a wide barrel syringe with a filter attached so that thesolution will be filtered immediately before use.
5) Preparing Subbed Slides. Heat 500 ml water to 80°C. Add 5g gelatin (Sigma) and 0.5g
chromium potassium sulfate. Stir on a hot plate until dissolved (about two hours).
Place new slides in racks, wash briefly in a warm hemosol solution, rinse with water.
Dip slides in gelatin solution, and dry covered overnight.

90

Richard Carthew

SOMATIC MOSAIC ANALYSIS IN THE EYESomatic clones in the eye can be generated by X ray-induced mitotic
recombination and can be genetically marked by the cell-autonomous white gene. For
any gene you want to analyze by generating clones, you want to use the marker whitegene at a cytological map position which is close to the gene of interest. This isconveniently done by choosing an appropriate P-element transformant stock in which
the P vector contains the white gene and has integated in the genome cytologically close
to the gene of interest, which will be called gene X. Three criteria should be considered
before choosing the transformant stock. One, the white gene should be locatedcyologically close to gene X. Two, it is preferable if the white gene is more proximal tothe centromere than gene X. Third, one copy of the white gene in this transformant stock
should produce enough pigment in the photoreceptor cells that it can be easily seen insemi-thin or thick sections. This last consideration can be rapidly surmised by looking
at flies hemizygous for the white gene under a dissecting microscope. Their eye color
should be indistinguishable from wild type. Alternatively, you could fix and section
eyes from hemizygous flies and score pigment in photoreceptor cells. However this last
criterion is most important when analyzing mosaic ommatidia and is not important if
you just want to look at the phenotype of the entire clone.
Make a fly stock which is homozygous for a mutant allele of gene X and which
is homozygous white minus. We use the null allele of white, w-1118. Obviously if themutant allele of gene X is lethal, it has to be balanced over a chromosome with adominant marker.
The strategy is to put the chromosome with mutant gene X in trans to thechromosome containing the white gene. Recombination events between sister
chromosomes during mitosis are induced by X irradiation. If recombination occurs
between the centromere and the white gene it will produce a daughter cell that ishomozygous mutant for white and gene X. Replication will expand cell numbers to thepoint that in the eye a white patch is seen in a red eye.
Make a mass fly cross between w-; X- and w-; P[w+]. I prefer to mate 400 of eachgenotype putting 100 pairs in one bottle. Incubate 24 hours at 25°C. Transfer flies tofresh bottles. These can either be egg laying bottles or regular bottles. I prefer egg laying
bottles because the parameters for irradiation are more consistent. If you use egg laying
bottles, use plates containing yeast-glucose medium (see end of section). Collectembryos for 12 hours at 25°C. Repeat collection. Incubate plates or bottles for a further
42 hours at 25°C. The flies will now be 48 +/-6 hours old and will be late first instar
larvae. Inducing recombination at this time gives a balance between reasonable clone
size and frequency of clones (about 1 in 20 flies). If you want to generate larger clones,
irradiate at an earlier time, but be prepared to generate clones at a lower frequency.
Irradiate tlarvae in plates or unstoppered bottles with 1000 rads (115 kV, 5 mA).
If using egg-laying plates, cut the plates in halves or quarters and place each piece in abottle. When the flies have eclosed, they can be screened for clones. Most white clones
appear black under a dissecting microscope. Fix and section the eyes as described in
Section 19. However, before the flies are sacrificed, shine a bright light on them for a
few minutes. This causes the pigment granules in the photoreceptor cells to migrate
apically and makes it easier to score for white pigment in cells R7 and R8. Cut 1 to 4

91

Somatic mosaic analysis in the eye

micron thick sections and collect all of the sections serially. This is important as pigment
may only be visible in R7 and R8 cells for one or a few sections. Do not stain the eye
sections but mount in DPX and view under phase contrast. Pigment granules appear as
black specks adjecant to each cell's rhabdomere.
Recipe for Yeast-Glucose Plates:
10% dry yeast
10% glucose
1.6% agar
3/100 (v/v) 10% tergocept
Boil water and add solids. Boil for 5 minutes while stirring. Pour into 15x60 mm
plates.

92

Bruce Kimmel

PREPARATION OF ADULT DROSOPHILA FOR SCANNING ELECTRONMICROSCOPYThis protocol describes two different methods for dehydrating adult Drosophila
in preparation for S.E.M. For S.E.M. the water in a specimen must be removed without
collapsing it by the surface tension forces which occur as the liquid evaporates. Air
drying is unsatisfactory for adult Drosophila. In addition to providing better
preservation, the two methods described below have the added advantage of rinsing theflies with an organic solvent and thus removing exterior dirt and grime. The first
method requires the use of a critical point drying apparatus which most E.M.
laboratories will have. The second protocol employs the use of a low surface tension
solvent such as Freon 113.
1) Anesthetize the adult flies with CO2 or your favorite anesthetic unless of course you
are a cruel and sadistic person.
2) Place ten flies into 5 mls of 25% ethanol and incubate for 12-24 hrs at r.t.
3) Further dehydrate the flies through 50%, 75% and 2 x 100% ethanol as in step 2
above. Once in 100% ethanol the exterior morphology of the fly eye is stable for atleast one month.
FURTHER STEPS FOR CRICTICAL POINT DRYING
4) Crictical point dry the samples. (Critical point drying is a process of exchanging theethanol for liquid CO2 under high pressure. Once the ethanol has been replaced withliquid CO2, the temperature of the critical point drier can be raised such that the CO2
becomes gaseous. The transition from liquid to gas at temperature X and pressure Y
is the critical point. The CO2 gas can then be vented to the atmosphere thus escaping
from the sample with very low surface tension and leaving the sample dry. Check
with your local E.M. person for details on how to safely use a critical point drier.
There is a good reason for calling these things bombs.)
FURTHER STEPS FOR LOW SURFACE TENSION SOLVENTS
4) Pass the samples through a graded series of Freon 113 in ethanol (25%, 50%, 75%, 2 x100%) with 12-24 hr incubations in each step of the series at r.t. After the last 100%
step, remove the Freon 113 and vacuum dry the samples. (Because of the effects
which fluorcarbons like Freon 113 have on the ozone layer you may want to use analternative low surface tension solvent. The solvent hexamethyldisilazane is reported
to be such an alternative solvent. The following reference claims that dehydration
through this solvent can give better results than critical point drying (J. Microscopy
121: 185-189).

93

Preparation of adult drosophila for scanning electron microscopy

COMMON STEPS
5) The dried samples can be stored indefinitely under vacuum with dessicant.
6) The samples are then mounted onto S.E.M. stubs using T.V. tube coat (Ted Pella Inc.).
This adhesive is nice because it provides a good black background for photography.
Care should be taken to mount the samples with the appropriate orientation so thatthe external structures of interest can be observed on the scope.
7) Sputter coat the samples with a 25-nm-thick coat of platinum.
8) The samples are now ready for viewing and should be stored over dessicant.

94

Nick Baker

PHOTOMICROGRAPHY
The purpose of this section is to collect together information about the types offilm used in the Rubin lab., with brief comments on their processing and uses.
Information on the use of compound microscopes in general, for photography orotherwise, is not included and can be found in publications such as the handbooks
published by the Royal Microscopical Society. Useful titles include:
Handbook #1"An introduction to the optical microscope", by S.
Bradbury, Oxford University Press, 1989 ISBN 0-19-856419-8.
Handbook #13"An introduction to photomicrography", by D. J. Thomson
and S. Bradbury, Oxford University Press, 1987 ISBN 0-19-856414-7.
Black-and-white negative film
Kodak Technical Pan is the film of choice for most purposes. Technical Pan canbe processed to give a wide range of contrasts according to need. When used at lower
contrasts a large range of tones are retained in the image; at higher contrasts small
differences are accentuated that pictorial films have more difficulty recording. Table 1
contains a set of developing conditions for Technical Pan at different contrast levels,and typical examples of their uses. When in doubt, lower contrast will preserve more
information and can be followed by printing on high contrast paper.
TABLE 1
Contrast Level

Speed
rating(ASA)

Developer

Typical Use

A Low(=pictorial)

25

Technidol LC1
15 mins (20°C)
11 mins (25°C)
8 mins (30°C)

B medium50

HC110, dil. F2
8 mins (20°C)

C high100

D very high150

HC110, dil. D2
6 mins. (20°C)
D19, undiluted
4mins (4°C)

pictorial applications, e.g.,
copying figures, photographing autoradiographs.
Also good for eye imaginaldiscs after DAB staining
most other uses, e.g.
sectioned material, in situs
or embryonic cuticle preps
in dark field
Polytene chromosomes3
embryonic cuticle preps. inphase contrast

A cautionary note: processing schedule 4. results in negatives of extremely highcontrast. Small gradations in lighting not visible to the eye will be recorded. Typicalproblems that can arise include diffraction patterns caused by dirt outside the plan of
1Technidol LC must be made from powder and used within 24 hours.
2 HC110 is stored as a working stock solution. Dilution D= 1 part working stock + 9 parts H 0.

2

Dilution F = 1 part working stock + 19 parts H20.
3 See use of monochromatic light below.

95

Photomicrography

focus (eg: on the top of the coverslip or undersurface of the slide), by air-bubbles inimmersion oil, or imperfect centering of the microscope light source. With short
exposures the time taken for the shutter to traverse the field may be enough to
produce an exposure gradient across the negative. It will be appreciated that only an
exceptionally well-maintained microscope can be used for such work.
Technical Pan is less useful when very fast exposure ratings are required, eg forphotographing fluorescence images. Instead use Kodak Tmax P3200. Processing
conditions for this film with HC110 are given in Table 2.

TABLE 2
HC110 dilution B
AB
C

Speed Rating (ASA)
1600
3200
6400

21°C
7.5
10
12

24°C
6
7.5
9.5

27°C
5
7
8

29°C
4.5
5.75
6.75

Use of monochromatic illumination with B&W film
The signal-to-noise ratio of black-and-white films can sometimes be increased byusing a monochromatic light source. This is because parts of the spectrum notabsorbed by the specimen (and which therefore contribute only "noise") are not used.
Also a narrow bandwidth should increase lens resolution. Examples are the use of agreen bandpass filter (550-600nm) to photograph orcein-stained chromosome
preparations, or a blue bandpass filter (450-500nm) to highlight pigment granules in
Drosophila eye sections.
Colour negative film
Colour negative films have some advantages over black-and -white. They
contain more information, and are developed and printed commercially both rapidly
and with little effort to yourself. We have preferred Fujicolour 100, but are trying the
new high-definition Kodak Ektar films. The main disadvantage is loss of control overthe printing process. Only the standard (low) contrast level is available, cropping andenlargement are expensive, and variation in colour balance and exposure are hard to
avoid. These last two points result from the strong red colour of the negatives, whichmust be filtered out by the printer. Inexpensive commercial labs invariably expose andbalance prints automatically with respect to some standard. Consequently there islittle point bracketing the exposure of your negatives or using colour filteredillumination, since the printer will likely correct all of this. Printing your own requires
equipment and skill.

96

Photomicrography

Colour slide film
Colour reversal (slide) films do not suffer these disadvantages, since what you
get back is the film you originally exposed, and the processing is invariant. Accurate
prints can readily be made from slides if required. The most used films are
Kodachrome and Kodak Ektachrome. These two have a different construction that
affects their properties. Kodachrome comprises three separate light-sensitive layers,
each of which is processed separately to generate one of the primary colours. Also a
non-reflective backing reduces halation. The resolution and colour-rendition of
Kodachrome are unparalleled. Also the thorough nature of the processing effectively
removes uncoupled dyes, increasing the stability of the image. In Ektachrome films all
the colour reactions occur within the same layer, limiting the accuracy of the colour
reproduction, but making processing easier. Ektachrome films are available in avariety of speeds. Also, Ektachrome slides are supposedly more stable in intense
illumination, such as found in a slide-projector. We tend to use Ektchrome 160ASA
routinely, because of its speed and ease of processing, Ektachrome P800/1600 for
fluorescent images, and Kodachrome only when the very highest standards are
required.
Daylight/Tungsten films
Ektachrome is availible balanced for daylight or tungsten lamp (3200K)
illumination. The colour balance of a lamp changes with its output. With an Axiophot
it is therefore best to use the tungsten light source always, and use the neutral-density
filters to adjust the light level. An 87A filter (which approximately corresponds to theblue microscope filter) converts tungsten illumination for use with daylight film.
"Professional" films
Both Kodachrome and Ektachrome are availible in a "Professional" grade that isslightly more expensive. These offer better quality control, more natural colour in thecase of Ektachrome, and more accurate speed rating. These films are sold and stored
refrigerated, and there is little point paying the extra unless you intend to do likewise.
Reciprocity
At low light levels the response of photographic films (like X-ray films) ceases tobe linear. The exposure time necessary increases out of proportion to the decrease inillumination, an effect called "reciprocity failure". Typically this becomes significant
for exposures >1sec. The axiophot control panel will correct for this effect if areciprocity factor, specific for each film, is entered. Some reciprocity factors
recommended by Zeiss are:
KODAK Technical Pan
3
EKTACHROME 160
5
EKTACHROME 400 (=P800/1600)
6
For colour films, however, reciprocity failure will be variable with different
wavelengths, a problem not completely overcome by these exposure compensations.

97

Kevin Moses

GENOMIC DNA PREPS
Large Scale:1) Freeze flies in liquid Nitrogen, in disposable polypropylene tubes, store at -70°C.
2) Set a porcelain mortar in dry ice, pour in liquid Nitrogen, and immerse the pestle until
cold.
3) Add frozen flies, grind as a slurry in liquid Nitrogen, then allow the Nitrogen to boil
off.
4) Chill a 50 ml polypropylene Oak Ridge tube and a spatula in liquid Nitrogen, thenadd up to 5 ml of the frozen fly powder to the cold tube on dry ice.
5) Remove the tube from the dry ice and immediately add 15 ml Homogenization Buffer(HB) and 15 ml 1:1 Phenol/Chloroform.
6) Cap the tube and mix by gentle inversion (to minimize DNA shearing) until well
mixed and then for 30 minutes on a lab-quake or similar.
WARNING - EXPLOSION HAZARD
AS THE FROZEN POWDER DISPERSES IT WILL RELEASE NITROGEN GAS. YOU
MUST PERIODICALLY CRACK THE CAP TO RELEASE THE PRESSURE AS THE
POWDER IS DISPERSED, UNTIL IT IS ALL DISSOLVED. OTHERWISE THE TUBE
MAY EXPLODE AND SPRAY NOXIOUS CHEMICALS ON YOU.
7) Spin at 18,000 rpm at 20°C for 10 minutes (if at 4°C the Urea and SDS in the HB will
precipitate).
8) Take the aqueous (top) phase, avoiding the interface, to a new tube and repeat theextraction twice more.
9) Take the aqueous (top) phase, avoiding the interface, to a new tube and add twovolumes of ethanol (don't add salt, it's already in the HB), mix and spin at 18,000 rpm
at 20°C for 10 minutes (if at 4°C the Urea and SDS in the HB will precipitate).
10) Resuspend the pellet in 3 ml TE by gentle inversion on a lab-quake or similar (thiswill take some time).
11) Add 3g CsCl and 0.3 ml 10mg/ml ethidium bromide, fill a 5ml ultracentrifuge tube
(top up with the same solution less the DNA pellet), check the density (ρ = 1.56) andspin at 45,000 rpm, 15°C for 16 hours.
12) Remove the DNA band by side-puncture (avoiding the pelleted RNA) and remove
the ethidium by repeated extraction with CsCl saturated butan-2-ol.
13) Dilute 3 fold with TE, add 1/10 volume 5M NaCl and precipitate with 2 volumes of
ethanol.

98Genomic DNA preps14) Wash the pellet in 70% ethanol, and resuspend in TE by gentle inversion.
15) Quantify by UV absorbance.
HB Buffer:
7M
2%
50 mM
10 mM
0.35 M

Urea
SDS
Tris pH 7.5
EDTA
NaCl

NotesThis protocol works by keeping the material frozen until it hits the combined HBand phenol/chloroform. Thus the cells "wake-up" into a chemical environment that
immediately denatures all membranes and proteins. In other words, the nucleases never
get a chance. This method works well as an RNA prep also, just vortex at each mixing
step, extract more exhaustively (until the interface is totally clean) and don't Caesium
band. The resulting total nucleic acids prep can be used to make polyA+ RNA on an
oligodT column, for Northerns, but if it is important to remove the DNA completeley (iefor cDNA cloning) then some measure must be taken to achieve this. This prep can bescaled down to 20 flies or fewer in a microfuge tube. For the mini-prep don't freeze, justhomogenize in 0.5 ml HB and then extract, twice with phenol/chloroform, ethanol
precipitate twice, wash in 70% ethanol and resuspend in TE. When you digest the miniprep DNA with restriction enzymes add boiled RNAse to 0.1 mg/ml.

99

Genomic DNA preps

David Bowtell

RNA ISOLATION USING GUANIDINE HYDROCHLORIDE
1) For adult flies that are frozen, first grind to a fine powder in a mortar and pestle
under liquid nitrogen. Use a large pestle to prevent "blasting" the powder out when
the nitrogen is added. Some of us have found that the addition of liquid nitrogen is
unnecessary.
2) Add the powder to room temp 6M GHCl, 0.1M NaOAc or 6M guanidine
isothiocyanate, 0.1M NaOAc. I generally use 10-20ml per scintillation vial of flies. It isbetter to add too much than too little liquid but first consider the number of samples
you are spinning in step 4. (A SW41 can take about 42ml of GHCl lysate per run).
NOTE. If you add the powder to a 50ml tube be careful shaking it up once the lid isscrewed on. Release of the dissolved nitrogen can blow the lid off. A beaker with theguanidine stirring in it is probably a safer alternative.
3) Shear the lysate through an 18ga. needle and then preclear twice at 10-15K in the SS34.
It is important to get rid of all the particulate material. The first preclear has a lot of
cuticle floating on the top.
4) Pretreat polyallomer tubes with 0.1M NaOH and rinse with DEPC DDW. Add 5.0ml
of 5.2M CsCl, 10mM EDTA and then layer the supernatant over the top. Spin (SW41,
SW40) 30K 16-18hr at 20° C.
5) Remove supernatant with a vacuum aspirator down to the last centimeter of liquid.
Often a prominant band is visible ~0.5cm above the pellet. This is (?) tRNA. The
pellet is usually not obvious until all the liquid is removed. Cut the tube off with ascalpel blade above the liquid and invert the tube. The pellet should be clear andgelatinous. Resuspend in ~200µl 10mM EDTA by shunting up and down with a p200.
It will take several minutes. Be careful not to let the suspension touch the end of thepipette. Remove 5-10µl into 400µl of 10mM EDTA for OD. Add the remainder toethanol/NaCl. It is best to store the RNA as a fine precipitate in ethanol at -70° C fromwhich aliquots can be removed.
6) To oligo-dT select, spin down RNA (very briefly if you have a lot otherwise the pellet
can be hard to resuspend). Resuspend in 10mM Tris, 1mM EDTA, 0.5M NaCl,
200µg/ml proteinase K and 0.1% SDS (it is not necessary to phenol extract) and either
oligo-dT select batchwise or over a column.
7) This works fine for adult flies, embryos, discs and probably all other stages.

100Marek Mlodzik

PLASMID RESCUE FROM P[lacZ] LINES
All of the commonly used P[lacZ] vectors contain sequences that allow plasmid
recovery (rescue) including genomic sequences from the transformed flies. Sequences
required for plamid rescue are a bacterial origin of replication and an antibiotic
resistance gene (e.g. Ampicillin or Kanamycin resistance). In addition, uniquerestriction sites are required that cut in the transformation vector only on one side of theori/resistance sequences; the second site is provided by flanking genomic DNA, so thatthe rescued plasmid contains DNA originating from both the transformation vector andthe fly's genome.
Depending on the vector, you can rescue genomic sequences upstream and/or
downstream of the P[lacZ] insert.
1) Extract DNA from 40 to 50 flies with your favourite procedure. An easy method is
described in the protocol for "Rapid small scale isolation of Drosophila DNA andRNA", p. 183 in this volume.
2) Check quality of DNA with desired restriction enzymes and Southern blotting.
Usually 2 to 4 fly equivalents are enough to see a "good smear" on agarose gel. This
step is optional, but K. (Mosey) Moses would not ommit it.
3) Digest 2 to 10 fly equivalents of DNA with an enzyme that fullfills desired criteria
for rescue (If several different sites are suitable, try with more than one.) If possible
use an enzyme that can be heat inactivated, otherwise include phenol/chloroform
extraction before precipitation of DNA after digest.
4) Resuspend digested DNA in 20 to 50 µl TE. Use half of the DNA for the ligation. To
avoid intermolecular ligation events, perform ligation reaction in a larger volume;
200 µl are usually fine. It is not necessary to add more than 2µl (e.g. 2 units of N.E.
Biolabs enzyme) T4 DNA ligase. Incubate reaction mix for 4 hrs to overnight.
5) To 200 µl ligation mix add 20 µl of 3 M sodium acetate and precipitate with twovolumes of ethanol. Spin, wash, dry and resuspend pellet in 10 to 20 µl TE. The DNA
is now ready for transformation.
6) For bacterial transformation, I have used XL1-blue cells (Stratagene) and a highefficiency transformation protocol as described by Hanahan (see below for details).
However, since more recently the electroporation procedure has been commonlyused and gives even higher transformation efficiency, this might be the method of
choice for plasmid rescue bacterial tranformation. When using electroporation, thecuvettes should be sterile to avoid "surprises". Plate cells on Ampicillin or
Kanamycin plates depending on transformation vector used.
Comments:
The number of colonies per fly equivalent of genomic DNA varies greatly,
ranging from 0 to 30. The use of XL1-blue cells (10 to 100 times greater transformation
efficiency than other commonly used bacterial strains) and 25 C/ Hanahantransformation protocol (see below) allows usually plasmid rescue from any
transformant line. Difficulties in obtaining clones from particular lines seems most likely
due to an unusual distribution of enzyme sites near the insert (try with different
enzymes if possible).

101

Plasmid rescue from P[lacZ] lines

When rescue of genomic upstream sequences is employed, the rescued plasmid
still contains the lacZ gene. For such plasmids, it is worthwhile to plate on plates
containg X-gal. Usually all XL1-blue colonies containing such plasmids will turn blue.
This might be usefull to destinguish the desired plasmids from potential contaminants.
High efficiency (Hanahan) competent cells
1) Pick a single XL1-blue colony to 5 ml LB, grow to saturation at 37°C.
2) Inoculate 50 ml 2xYT + 10mM MgCl2 and 0.1 % glucose with 100ml of the seed culture in a
250 ml flask and grow at 25°C to A600 = 0.3 (about 12 hr).
3) Spin down cells in a 50 ml orange cap tube, resuspend cells in 20 ml ice cold TFB andincubate on ice for 15 min.
4) Spin down cells and resuspend in 4 ml TFB.
5) Add 140 ml DnD and incubate on ice for 10 min.
6) Add 140 ml DnD, aliquot 200 ml per transformation and incubate on ice for 20 min.
7) Add up to 20 ml DNA and incubate on ice for 20 to 40 min.
8) Heat shock at 42°C for 90 seconds then return to ice for two min.
9) Add 800 ml 2xYT (+ 10mM MgCl2 and 0.1 % glucose), incubate at 37°C for one hour.
10) Plate, (if M13 phage wash the cells with two changes of medium first)
Solutions for Hanahan Cells:
TFB (for 500 ml)
make 100 ml 0.5 M MES pH 6.3(9.76 g MES, pH with KOH, filter sterilise)
3.70 g Ultrapure KCl
4.45 g MnCl.4H2O
0.75 g CaCl2.2H2O
0.40 g Hexamine CoCl3
10 ml 0.5 M K-MES pH 6.3
Distilled Water to 500 ml, filter sterilise, store at 4°C.
DnD

Reference:

1.53 g DTT
9 ml DMSO
100 ml 1 M Potassium Acetate (pH 7.5)
aliquot and store at -70°C.
Hanahan, D., Techniques for transformation of E. coli
Ch 6 in DNA cloning, Vol I, A practical approach
Ed Glover, D.M., Oxford, IRL Press

102

FINAL
100 mM
45 mM
10 mM
3 mM
10 mM
1 M
90 %(v/v)
10 mM

Ulrike Heberlein

PREPARATION OF PROBES FOR FOOTPRINTING EXPERIMENTS
Probes are essentially labeled as for Maxam and Gilbert sequencing; one end of
the DNA fragment is end-labeled with 32P-γATP. The general strategy is to cut theplasmid with a restriction enzyme at the site to be labeled. The 5'-phosphate is removed
with alkaline phosphatase and replaced with 32P. The labeled DNA is then cut with asecond restriction enzyme and the fragment of interest is isolated. It is important tomake sure that a combination of digests will produce fragments that can be separated
from each other by gel electrophoresis. Restriciton sites that result in a 5' overhang
should be used; however, blunt ends can also be labeled with lower efficiency.
1)

First digest: Digest ~5-10µg of CsCl-purified DNA (any RNA contamination will
greatly reduce the efficiency of the labelling!!) with the restriction enzyme of choicein a 50µl volume. Check the digest on a gel to make sure it is complete (partial
digests can cause nightmares).

2)

Removal of 5' phosphate: To the above digest add: 45µl of H2O + 2.5µl of 2M
Tris/HCl pH9.5 + 2.5U of calf intestine alkaline phosphatase (from Boehringer
Mannheim, 1U/µl, cat. #713023). Incubate at 37°C for 30-45 min. Extract with 1
volume of 1:1 phenol/chloroform twice (this is important to get rid of all thephosphatase before labelling). Extract with 1 volume of chloroform and ethanol
precipitate with 2.5 volumes of ethanol in the presence of 0.3M NaOAc.
Resuspend the washed and dried pellet in 16µl H2O (this volume is based on thevolumes and concentrations of reagents used in the kinasing reaction described
below; adjust it if you need to).

3)

Addition of 32phosphate: To the phosphatased DNA in a 16µl volume add: 2µl
10X kinase buffer (0.5M Tris pH 8, 0.1M MgCl2, 50mM DTT, 1mM EDTA pH 8,
10mM spermidine; store at -20°C) + ~1µl 32P-γATP (~150µCi, ICN crude,
7000Ci/mmole) + 1µl polynucleotide kinase (10U/µl). Incubate at 37°C for 30-45
min. followed by 15 min. at 65°C to kill the enzyme.

4)Second digest: To the labeled DNA add: 5µl of 10X restriction enzyme buffer, 1020U of the appropriate enzyme, and H2O to a final volume of 50µl and incubate at
37°C for 1-2 hrs. If you want to reduce the volume of radioactive liquid waste,
you can precipitate the digest at this point to eliminate the non-incorporated 32PγATP.

5)

Isolation of labeled fragments: Separate the labeled fragments by gelelectrophoresis. If you need to resolve small fragments use a 5% or 7% nativeacrylamide gel. Otherwise use a regular agarose gel (TAE buffer). Remember thatthese gels a screaming hot!!

103

Preparation of probes for footprinting experiments

A)

Acrylamide gels: ~20cm long gels, 1.5mm thick are adequate. Add sucrose
dye to the DNA, heat at 65°C for 5 min., load onto the gel and run for 2-3
hrs at ~200V. When finished, take off the top plate and cover the gel
together with the bottom plate with Saran Wrap. Expose to film for 1-2
min. The wells are usually visible on the autradiogram, allowing easyalignment of film and gel; however you may want to use radioactive
markers. Cut out the band of interest and electroelute in 0.5X TBE for 30-60
min. Precipitate the eluate with 1 volume of isopropanol at -20°C for 1-2
hrs. Spin for 15 min in a microfuge at ~15,000RPM. Resuspend the DNA in
50-100µl of TE and store at -20°C. About 0.1-0.5µl of this DNA should beenough for one footprinting reaction. If the concentration of DNA-binding
factor to be assayed is low (as in most cell or tissue extracts) the minimum
amount of probe, hopefully of high specific activity, necessary to be
detected in an overnight exposure should be used. 1µl of labeled probeshould give 50-200 cps on a minimonitor.

B)

Agarose gels: Run a 0.8-1.2% TAE/agarose gel, stain with ethidium
bromide, cut out desired band and isolate the DNA using Gene-Clean. (Ipresume that low melt agarose would also work, but I have never tried it.)
Resuspend and store as above.

Purine cleavage sequence markers(for footprinting experiments):•Take 8µl of footprinting probe (~80,000 cpm Cerenkov) labelled as described.
•Add 5µl of 1mg/ml sonicated calf thymus DNA and dry in the speed-vac.
•Add 6µl of 2% formic acid (diluted fresh from 88% Malinckrodt stock stored at 4°C)
•Incubate at 37°C for 12 minutes and put on ice immediatly. This time is optimal toobtain an evenly cleaved ladder (~1 hit per molecule) of a 300bp long probe.
•Speed-vac for 2 hrs.
•Add 100µl of a 1:10 dilution of piperidine (Sigma, prepared fresh in the hood!!).
•Incubate at 92°C for 20 minutes. Put a weight on top of the tube(s) or they will pop
open and release very toxic piperidine.
•Speed-vac overnight.
•Add 50µl ddH2O, dissolve and speed-vac until dry.
•Resuspend in 20µl of 0.1N NaOH + 40µl formamide dye.
•Use 1-5µl for each lane.

104

Ulrike Heberlein

DNAaseI FOOTPRINTINGThis technique exploits the fact that a protein bound to a specific DNA sequence
will interfere with the digestion of that region by the endonuclease DNAaseI. In
summary, an end-labelled DNA probe is incubated with a protein extract or a purified
DNA-binding factor. The unprotected DNA is then partially digested with DNAaseI
such that on average every DNA molecule is cut once. Digestion products are thenresolved by electrophoresis. Comparison of the DNAaseI digestion pattern in thepresence and absence of protein will allow the identification of a footprint (protected
region) or hypersensitive sites (sites at which digestion is enhanced due to protein
binding).
Binding reaction:
Mix in a 50µl final volume:
0-25µl protein extract in HEMG (see below)
10µl
10% polyvinyl alcohol (store 4°C)
1µl
1M Hepes pH 7.6 (pHed with KOH)
~2fmoles of end-labelled DNA probe (~20cps on minimonitor)
competitor DNA (see below)
ddH2O to a final volume of 50µl
If the amount of protein used is less than 25µl, add 0.1M HEMG to make up the
difference. Mix components gently and incubate ~10 minutes. Incubation on ice isusually required when dealing with relatively crude protein extracts which containendogenous nucleases and/or phosphatases. With purified proteins the incubation
temperature can be increased.
It is advisable to add non-specific competitor DNA, such as sonicated calf
thymus DNA, synthetic poly d(I-C) or poly d(A-T) (Pharmacia) to the binding reaction.
The optimal competitor concentration needs to be determined empirically, however 0.10.5µg of sonicated calf thymus DNA or 0.01 OD units of synthetic DNA are usuallyadequate.

105

DNAaseI footprinting

DNAaseI digestion:
To the binding reaction add 50µl Ca/Mg solution (10mM MgCl2, 5mM CaCl2) atroom temperature and 1-10µl of DNAaseI solution (see below). Mix quickly andincubate at room temperature for 1 minute. Stop the reaction by adding 100µl of stop
solution (0.2M NaCl, 20mM EDTA pH 8, 1% SDS, 0.25 mg/ml carrier RNA; store atroom temperature) and vortexing immediately. Digestion of three samples can be
carried out simultaneously with a bit of practice.
Samples are then extracted with 200µl of 1:1 phenol:chlorophorm andprecipitated with 1ml of 100% ethanol. Pellet is washed with 70% ethanol, dried andresuspended in 6µl of formamide dye. Run on 6-8% sequencing gel.
HEMG:

0.1M KCl
25mM Hepes pH 7.6 (pHed with KOH)
0.1 mM EDTA pH 8
12.5mM MgCl2
10% glycerol
1mM DTT (fresh)

DNAaseI: Make a 2.5mg/ml DNAaseI (Worthington or Cooper Biomedical, bovinepancreas, DPFF, 2000U/mg) solution in ddH2O and freeze 2-5µl aliquots; keep at -70°C.
Make a fresh dilution of this stock solution in ice cold ddH2O and keep on ice. For the
no-protein control use ~1µl of a 1/1000 dilution. For crude protein extracts the amountand dilution of DNAaseI needs to be determined; 5µl of a 1/100 dilution are usuallyadequate.

106

DNAaseI footprinting

Ulrike Heberlein
PRIMER LABELLING FOR PRIMER EXTENSION ASSAY
Mix in a 20µl final volume:2µl 10x kinase buffer
1µl oligonucleotide primer (5pmoles/µl)
200µCi 32P-γATP (crude, ICN, 7000Ci/mmole)
10 U polynucleotide kinase
ddH2O to 20µl
Incubate at 37°C for 30-45 minutes. Heat at 65°C for 15 minutes.
To remove unincorporated label precipitate as follows:
Add 60µl TE + 320µl 2.5M NH4OAc + 20µg RNA carrier + 650µl 100% ethanol (follow
this order of addition; don't add the carrier RNA to the reaction first or some RNA will
get labelled). Chill on dry ice for 5-10 minutes. Spin in the cold for 15 minutes. Discard
supernatant (very hot!!). Resuspend pellet in 180µl TE and add 20µl 3M NaOAc + 500µl
100% ethanol. Chill on dry ice for 15 minutes, spin in the cold for 10 minutes, wash with
70% ethanol, dry and resuspend in 100µl TE (should be ~200cps/µl on minimonitor).
10x kinase buffer:
500mM Tris pH 8
100 mM MgCl2
10mM spermidine
1 mM EDTA
50mM DTT (add fresh before using)107

DNAaseI footprinting

Ulrike Heberlein

PRIMER EXTENSION
All solutions, eppendorfs and pipet tips need to be RNAase-free and sterile. Usegloves and work carefully (don't sneeze in your tubes).
1)

Precipitate the RNA with 5' end-labelled primer in 0.3M NaOAc with 2.5
volumes of ethanol. If working with low amounts of RNA add some yeast RNA

2)carrier (approx. 10µg). Wash pellet with 70% ethanol and dry.
Resuspend RNA in 8 µl ddH2O and add 2 µl of 5X annealing buffer (1.25M KCl,
10mM Tris-HCl pH 7.9, 1mM EDTA). Double this volume if more than 20 µg of
RNA are used, i.e., use 16 µl ddH2O and 4 µl of 5X buffer.

3)

Incubate at 60° for 90 minutes. Spin briefly every 30 minutes to avoid drying of
the RNA. The annealing temperature depends on the length and GC content of4)

the primer. The above is O.K. for 20-30 base primers with 50-70% GC content.
Add 23 µl of PE mix (10mM MgCl2, 5mM DTT, 100 µg/ml actinomycin D,
0.33mM dNTP's, 20mM Tris pH 8.7 at room temp., pH 8.3 at 37°C, store at -20°C
in the dark) and 10 U of reverse transcriptase (Life Sciences). Mix carefuly, spin
briefly, mix again. (Use twice these quantities if annealing was done in 20 µl.)5)

Incubate at 37°C for 1 hour.

6)

Add 0.3 ml chilled ethanol. Put in dry ice bath for 15 min. Spin at 4°C for 10
min. Wash in 70% ethanol. Dry.

7)

Dissolve in 2 µl of 0.1N NaOH, then add 4 µl of formamide dyes. Boil for 2 min.,chill on ice and load on sequencing gel.

108

Matthew Freeman

OLIGO-LABELLED DNA PROBESThis technique is an alternative to nick translation, and is generally superior. I findit less fussy, and it routinely gives considerably hotter probes. It also has the advantagethat even quite dirty DNA preps work fine, and it is possible to label fragments in LGT
agarose with no purification.
The 5X oligo-labelling buffer (OLB) can be made in large batches and stored at -20° for atleast a year. If you do this, keep it in aliquots of a size such that you thaw and refreeze aparticular tube for not more than a month or two - after this it may become less effective.
For all normal uses it is only necessary to use one hot nucleotide (I use 32P-dATP).
The OLB has the necessary cold nucleotides, so you need to make another one if you
want to use two hot ones.
The OLB is made from several component solutions.
Solution O:

1.25M Tris HCl, pH8.0
1.25M MgCl2

Solution A:

2mls Solution O
36µl ß-mercaptoethanol
10µl 100mM dATP
10µl 100mM dGTP
10µl 100mM dTTP

Solution B:
TE

Random hexamers (Pharmacia), 90 OD Units/ml in

10mls 5X OLB:

2mls Solution A
5mls 2M HEPES, pH6.6
3mls Solution B.

109

Ologo-labelled DNA probes

DNA labelling:
1) Boil DNA in 32µl (made up with water) for 5-10 minutes, then incubate the DNA at
37° for 5 minutes (this is clearly black magic, and may not be necessary).
2) Set up the reaction as follows: DNA

32µl
5X OLB
10µl
10mg/ml BSA
2µl
32P-dATP(3000 Ci/mmol)
5µl
Klenow
1µl (2-5U)
The reaction is done at room temperature; most incorporation has ocurred by one
hour, but it continues to increase till about 3-4 hours. Unlike nick translation, it does
not then diminish, and if it's convenient, it's fine to leave it overnight.

3) Separate unincorporated nucleotides by your favourite method: mine is small spincolumns in 0.5ml Eppendorf tubes.
4) You will find routine incorporation of between 50% and 80%, unless you used lessthan 20-30ng of DNA. The specific activity can reach about 3x109 dpm/µg.
To label a DNA fragment in LGT agarose substitute the following for step 1 above.
1a.)Cut the band from an LGT gel and add 3mls of water for every gram of agarose(Eppendorf tubes weigh 1g). Boil for 10-15 minutes, then place at 37° for 10 minutes.
Use 32µl of this mix in the normal reaction. The remaining DNA/gel mix can befrozen and reused many times (I boil it each time, but am not sure that it'snecessary).
Notes:a) The quantity of DNA: anything over about 5ng works; theoretically 20-40ng gives thehottest probe in the above reaction; over 100ng will reduce the final specific activity.
b) I have found one or two cases (out of many) where a supercoiled plasmid doesn'tlabel well. This can be helped by a longer boiling step or, in desperation, linearising
the plasmid before labelling.
Refs:
A.P. Feinberg and B. Vogelstein.

Anal. Biochem. 132, 6-13 (1983)
Anal. Biochem. 137, 266-277 (1984)110

Matthew Freeman

NORTHERN BLOTS USING RIBOPROBES
The advantage of using riboprobes for northerns is that you can make very hotprobes, and the extra stability of RNA:RNA hybrids may increase the sensitivity. They
are therefore useful if you are looking for similar, but not identical, transcripts, or if yourtranscript's abundance is very low. There are significant disadvantages also. In orderto make the hottest possible probe, a relatively large amount of radioactivity is used, andalso there can be a problem with spurious bands (for example I often see the same band
when I have used Bluescript vectors as the probe, presumably because there is a degreeof similarity between some vector sequences and some Drosophila transcript). The latterproblem is usually solved by increasing the stringency of the final washes: the different
signals melt off very efficiently at different temperatures, and you could normally expectthat the most stable hybrid represents the 'real' transcript. Nevertheless, this could
clearly cause problems in some situations, as well as actually being an advantage inothers (such as deciding which of several transcripts was most similar to a given probe).
It should be noted that these probes are RNA, and are therefore sensitive to RNAses,
which contaminate most glassware, and all hands. Use normal precautions for working
with RNA and all will be well. The protocol below takes a very cautious approach; inpractice you can usually be more sloppy.
Labelling the Probe:
The following recipe gives the hottest probe that can be made from available
nucleotides. It uses 750µCi to make a single probe, and therefore should only be used in
extreme situations. You can reduce the amount used by using ribonucleotides of lower
specific activity and/or reducing the reaction volume; the critical factor is that theconcentration of the hot nucleotide is at least 8µM. The whole procedure is based on the
one in the instructions that come with the Stratagene Transcription Kit; there is a
discussion of all the stages in that booklet.

111

Northern blots using riboprobes

1) Dry down 750µl of 3000Ci/mmol, 1mCi/ml rATP in the Speedvac (I split it into two
tubes, thereby halving the time taken).
2) Resuspend the hot nucleotides in the following reaction:
5µl 5X Transcription Buffer* (Stratagene)
1µl 10mM rCTP
1µl 10mM rGTP
1µl 10mM rUTP
1µl 0.75M DTT
1µl RNAse Block (25U, Stratagene; RNAsin is OK too)
1µg DNA†
10U T3 or T7 Polymerase
Water to 20µl
*5X - 200mM TrisHCl, pH8, 40mM MgCl2, 10mM spermidine, 250mM NaCl.
† The DNA is Bluescript cut down stream of the insert, treated with 200µg/ml
proteinase K, 15 minutes, 37°, phenol extracted, ethanol precipitated, andresuspended in DEPC-treated water.
3) Incubate at 37° for 30 minutes.
4) Add 100µg RNAse-free tRNA, an equal volume of 4M ammonium acetate, and 2.5
volumes ot ethanol. Precipitate. By measuring the radioactivity in the pellet and thesupernatant, it is possible to get a very approximate idea of the incorporation; itshould be greater than 50%. Resuspend in boiling salmon sperm DNA just before
setting up the hybridisation.
Northern Blot
The northern is run under standard conditions.
Hybridisation
1.Prehybridise for at least 4 hours at 65° in:
50% formamide
5X SSC
1X PE*
150µg/ml salmon sperm DNA (phenol extracted)*5X PE is:

250mM Tris-HCl, pH7.5
0.5M sodium pyrophosphate
5% SDS
1% polyvinyl pyrolidine
1% ficoll
25mM EDTA
Make in DEPC-treated water, heat to 65° to dissolve. Cool to about 37°, and add 5%
BSA (fraction 5) to 1% final. Heat again to 65° for 15 minutes. Store indefinitely atroom temperature.

2) Hybridise in the same mix with the probe (I resuspend the probe in a little just-boiled
salmon sperm DNA). Hybridise at 65°, overnight or longer.
3) Wash in 0.1X SSC, 0.1% SDS at 65°. This is rather low stringency (see above), and any
signal you see should be checked by rewashing the filter (which should not be112Northern blots using riboprobes

allowed to dry completely) at increasing temperatures. For example I have foundthree bands on a northern that only gives one with DNA probes. One of the spurious
bands comes off between 65° and 70°, and the other by 75°. In this case the real bandstill hybridises up to 85°.
Using this protocol, a transcript that is normally visible in about 12 hours (using oligolabelled probes), is visible in 1 - 2 minutes, suggesting that these really are very sensitive
northerns.

113

Smadar Admon

POLYCLONAL AND MONOCLONAL ANTIBODIESPOLYCLONAL ANTIBODY
Immunization of Mice:
Injections should be made at intervals of at least two weeks.. We have beenusing either of the two following adjuvants with success: Freund adjuvant and
MPL+TDM adjuvant ( RIBI ImmunoChem Research, Inc. ), the latter being lesshazardous than the first.
1) 1st injection: 15 µg Antigen in 200 µl adjuvant, injected subcutaneously.
2) 2nd injection: 2 weeks later; boost with 15 µg of Antigen in adjuvant. (If time
permits, boost again a month later).
3) 7 - 10 days later bleed the mouse and test serum using the assay which will be used
for screening. If titre is not high enough, boost again two weeks after previous
boost.

Serum Preparation:1) After collecting, blood should be allowed to clot for 60 min . at 37°C or O.N at 4°C.
2) Separate the clot from the sides of the tube (ringing) using a pasteur pipette. Placeclot at 4°C O.N.
3) Spin at 10000 x g for 10 min. at 4°C to separate the serum.
4) Serum can be stored at -20°C after adding Glycerol to 50%.
If monoclonal antibodies are desired, once you have a "good" polyclonal and a reliable
screening method proceed to the next step.

114

Polyclonal and monoclonal antibodies

MONOCLONAL ANTIBODY
If titre is positive do one of the following 2 weeks later:
I- Boost with 50 µg - 100 µg of Antigen in PBS and proceed with fusion 4 days
later.
OR
II- Boost 3 days in a row with 15 µg Ag in PBS and proceed with fusion on 4th day.

Cell Lines:
1) Myeloma P3X63-Ag8.653
Origin: BALB/c, non secreting, 8-azaguanine resistant, HPRT -.
2) Myeloma fox-NY
Origin: Robertsonian, 8-azaguanine resistant, HPRT -, APRT -.
(mice have resistance to drug and expression of heavy chain on the same
chromosome).
3) Macrophage-derived J774A.1
Maintenance of the cells:
Stock solutions:•IMDM: JRH Scientific 51-47178.
•FCS: Fetal Bovine Serum- defined, Hyclone A-1111-L (no heat-inactivation needed)or any other brand that has been sampled and checked in advance.
•Insulin: 100X stock= 20 I.U./ml
Dissolve 80 mg of Insulin (25 I.U./mg) in 90 ml of ddH2O by adjusting to pH 3.0
with 1-2 drops of 6N HCl. After Insulin has dissolved readjust pH to 7.2 with 0.5
M Na2HPO4, bring volume to 100 ml and filter sterilize with 0.2 µ membrane,keep frozen.
•Oxaloacetic Acid: 100X stock = 0.1 M
Filter sterilize, keep frozen.
•D(+) Glucose: 100X stock = 45%
Filter sterilize, keep frozen.
•Transferrin: 1000X stock = 1 mg/ml
Dissolve 1 mg human transferrin for each ml of IMDM. Make a 2 mg/ml
FeCl3.6H2O or 1.2 mg/ml FeCl3 in 0.001 M HCl. Add 1µl Fe solution to each
ml of transferrin solution. Mix, let stand on ice for 30 min. Filter sterilize, store
at 4°C.
•HT supps: 100X stock = 1.36 mg/ml (H) , 0.73 mg/ml (T).
Weight out 68 mg hypoxanthine ( Sigma H-9377) and 36 mg of thymidine (Sigma
T-9250 ), combine. Add 1N NaOH (a few drops to 1 ml) and stir until the115Polyclonal and monoclonal antibodies

powder is in solution. Bring to a final volume of 50 ml with ddH2O. Filter
sterilize, store at -20°C.
•Supplements: 25X stock (store at -20°C).
Mix equal volumes of the following:
L- Glutamine (200mM = 100X) , Gibco 320-5030.
Penn-Strep (100X), Gibco 600-5070PG.
Non-essential amino acids (10 mM = 100X), GIBCO 320-11404G.
Sodium pyruvate (100 mM = 100X), Sigma S 8636.
•2-Mercaptoethanol: 1000X stock (5 x 10 -2). (Store at 4°C.)•AT supplement : 50X stock , Sigma A-7422. (Store at -20°C.)•PCM: Parents Conditioned Medium.
Medium is collected from the the apropriate parent myeloma after the cells havereached high density, then spun and filtered. Aliquot and keep at -20°C.
•MCM: Macrophage Conditioned Medium. (used instead of feeder cells)
Seed macrophages at a density of 1.5x105 cells/ml in the medium described on
next page. Add 2.5 µg/ml LPS which induces differentiation.
Collect sup after 2-3 days, or when medium is getting too yellow. Induce 2 moretimes , each time with 1 µg/ml LPS and collect sup after 2 days each.
Pool the sups, filter and use as recomended. (Could be aliquoted and stored at-20C.)
Preparation of Media:
for P3X63-Ag8.653: IMDM complete
to 400 ml of IMDM add:
5 ml 100X Insulin
5 ml 100X Oxaloacetic acid
5 ml 100X D(+) glucose
0.5 ml 1000X transferrin
0.5 ml 1000X 2- Mercaptoethanol
5 ml 100X HT
20 ml 25X supplements
75 ml FCS ( final 15% )100 ml PCM ( Parent conditioned medium ).

116

Polyclonal and monoclonal antibodies

for fox-NY: IMDM or RPMI (JRH Scientific, 51-50178).
10 % FCS
1X AT supplementtransferrin
supplements
PCM, when available from this cell line .
for J774A.1: out of convenience we have been using the same medium as the one for
the P3X63-Ag8.653 ( = Ag8 ), without the PCM.
Growth conditions:
All cell lines mentioned above grow at 37°C, 7% CO2 .
5
The Myelomas optimal density is 3.5x10 /ml.
The Macrophages optimal density is 1.5x105/ml. When expanding them , use a"policeman" to scrape them; this is easier to do if they are growing in petri dishes atthis stage.
Freezing Hybridoma / Myeloma / macrophages.
Freezing solution: 90% FCS + 10% DMSO, ice cold.
1) Spin down 107 cells (106 minimum) at 1200 rpm for 5 min.
2) Aspirate medium.
3) Resuspend in 1 ml of ice cold freezing solution.
4) Transfer vial to an insulated freezing box and place at -70°C for at least 1 hr. (could
be for a couple of days).
5) Transfer the vial from the -70°C to the liquid nitrogen tank and log the entry in thefreezer log.
Thawing cells:1) Take vial out of liquid nitrogen tank and thaw it immediately in a 37°C bath (about
1 min).
2) When there is still a small piece of ice left, dilute the cells by transfering them into a
conical tube containing 10 ml of the growth medium at 37°C.
3) Spin at 1200 rpm for 5 min.
4) Aspirate medium and resuspend cells in 5 ml of medium, in a 25cm2 flask.

117

Polyclonal and monoclonal antibodies

Cell Fusion and Selection
Solutions:•IMDM•IMDM complete with MCM instead of PCM.
•PBS
•PEG 50% ( w/v ): ( PEG 4000 "gas chromatography grade" Art.9727 - Merck). Soldby Bryant Laboratories, (415)526-3141, or MC/B Manufacturing Chemists, Inc.
•Aminopterin: Sigma, A 5159 (50X ), use it as 100X. ( final = 2x10-7 M ).
1)

Prepare a T-150 flask with 170ml of IMDM complete with MCM instead of PCM(= IMDM - m ), and keep it in the incubator for fused cells.

2)

Isolation of spleen cells :a ) Sacrifice mouse by cervical dislocation. Immerse mouse in 70% Ethanol.
b ) Remove spleen ( on the left side ) and transfer into a small petri dish whichcontains IMDM at room temp. Clean fat from the spleen and transfer the
spleen into an empty dish.
c ) Using sharp tipped forceps, one end is punctured. A curved forceps is used tohold down the intact end, and the spleen is gently rubbed towards the opened
end with another set of forceps. The cells from inside the spleen will ooze out
with very little damage. Stop the process when you are left with a nearly
empty, transparent skin. Collect the cells by rinsing with IMDM.
d) Transfer cell suspension to a 15ml conical tube and let the cell debris settle out(approx. 5 min.).
e) Remove the cell suspension (without disturbing the settled cell debris) andtransfer to a 50ml conical tube. Add an additional 30ml of IMDM and pellet
the cells at 1200 rpm for 5 - 10 min.
f) Aspirate the medium, resuspend pellet and wash again with 30ml of IMDM.
One immunized spleen has aprox. 108 cells. After this wash the cells areready for the fusion.

3)

Myeloma cell preparation:
It is essential that the myelomas be free of debris, rounded and refractive underphase contrast, and that they are harvested in log or late log phase growth(between 3.5 and 9 x105 cells/ml).
a) Thaw cells 7 days before scheduled fusion. Myeloma do not grow well after
being in culture for more than a few weeks.
b) Make sure you refreeze cells for future use.
c) We have been using a ratio of 2 spleen cells : 1 myeloma cell. However,workers have been using a ratio from 1 to 10 spleen cells per myeloma
successfuly. For one spleen we harvest 5x107 cells. It is advisable to do thisspin at the same time that the second wash of the spleen cells is done.

4)

The fusion:

118

Polyclonal and monoclonal antibodiesa) The washed myeloma and spleen cells are pooled in 30ml of PBS (room temp.)
and spun gently at 1000 rpm for 10 min.
b) Aspirate the PBS and resuspend pellet gently by tapping the tube. Volume
should be approx. 0.8 ml.
c) Set a timer.
d) Add an equal volume of PEG solution, slowly, dropwise, with gentle tapping,over 1.5 min. at room temp. Then gently wiggle the tube for 1.5 min.at 37°C.
Some cell clumping will be evident.
e) The suspension is spun at 1000 rpm for 3-4 min. (at this point you should see
the different layers of cells with PEG on top).
f) Slowly add 37°C IMDM to 10 ml, without disturbing pellet. After adding ,swirl the tube gently to mix and dilute the PEG. Do not disturb the cells.
g) Spin at 1000 rpm for 5 min.
h) Aspirate medium, resuspend cells by tapping. Slowly add 5 ml of 37°C
IMDM-m.
i) Bring to 20 ml and add to the flask in the incubator. (If using feeder cells, add
them at this point; 106 cells/ml).
j) Add Aminopterin (2ml to 200 ml of medium). (Some workers will leave thecells at this point for 24 hours before adding Aminopterin. We add the drug
immediately.)k) Seed the cells in 96 well microtiter dishes, 250 µl per well, 8 plates per fusion.
First clones may be seen in 7-10 days. First screen will usually start after 2 weeks,with a second and third, if necessary, a few days later.

Screening
ELISA
Materials:• Plates - 96 well Dynatech Immulon, type 2. (Fisher 17-0221-199).
• Adsorbtion buffer 10X = 1M NaHCO3 pH 9.6 ( 80ml 2M Na2CO3 + 170ml 2M
NaHCO3 + 250ml ddH2O ) or PBS.
• PBS, TBS• PBS + 0.1% Tween 20 or TBS + 0.1% Tween 20.
• Blocking solution = 2% BSA (type V ) in PBS. (Add 0.02% azide for longer storage.)• Elisa buffer = 2% BSA + 0.1% Tween 20 in PBS ( azide optional ).
•Diethanolamine Ph 9.8 ( 48.5 ml diethanolamine + 0.1 gm Na Azide + 0.05 gm
MgCl2.6H2O in 400 ml ddH2O. Adjust pH to 9..8 with concentrated HCl. Bring to
500 ml., store at 4°C in dark bottle).
• Enzyme linked antibody = Alkaline phosphatase Goat anti mouse. ( Zymed 62-6422). 1• Substrate = Sigma 104 phosphotase substrate tablets ( Sigma 104 -105 ).1
Protocol:1 This is the system we have been using mostly, but there are other linked antibodies

available which can be used with the same procedure.

119

Polyclonal and monoclonal antibodies

1) ADSORBTION OF ANTIGENa) Dilute Ag to 10 µg/ml in 1X adsorbtion buffer.
b) Add 100 µl of Ag solution to each well. (A good control would be to leave an
empty well with no Ag for every well that contains it.)c) Leave O.N covered with saran wrap , at 4°C.
2) BLOCKINGa) Wash unbound Ag by inverting the plates and flicking the wells dry.
b) Rinse by adding PBS to each well and inverting it again (use squirt bottle).
c) Repeat the rinse twice.
d) Add 100 µl of blocking solution to every well, leave 1 hr at room Temp or O.N
at 4°C.
3) PRIMARY ANTIBODYa) Add the antibody to be tested :Sup of cells = 25 µl , mix well by pipeting up and down (10 times).
serum, ascites = 1:500 to 1:1000 dilution. Do dilutions in blocking solution.
b) Leave 1 hr at room temp or O.N at 4°C.
4) SECONDARY ANTIBODYa) Wash unbound antibody 4 times with PBS + 0.1% Tween 20.
b) Add 100 µl of enzyme linked antibody to all wells. Do the appropriate
dilutions in the Elisa buffer. (ex: AP is 1000X).
c) Leave 1 hr at room temp or O.N at 4°C.
5) SUBSTRATEa) Dissolve substrate in Diethanolamine (for AP substrate, warm 37°C, 5 min.)to concentration of 1 mg/ml.
b) Wash plate 4 times with PBS + 0.1% Tween 20 (use TBS instead of PBS for AP).
c) Add 100 µl of substrate to every well.
d) Watch color development. This could take from a few seconds to 20 min.
e) If needed, stop the reaction by adding 50 µl of 4M NaOH.
f) Read absorption in Elisa reader at correct wavelength (for AP system 405 nm).

120

Polyclonal and monoclonal antibodies

Expanding Hybridoma Clones
1) Transfer the srongest positive clones to a 24 well plate by scraping the adhering
cells with a truncated yellow tip.
2) Bring the volume to 1 ml using the IMDM-m medium.
3) When cells reach confluency, freeze them ( about 106 cells per full well ). Scrapethem using a blue tip.
4) Add 1 ml of IMDM-m to cells left in the well, let them grow and repeat the freezing.
5) While the cells are growing, save their sup at each freezing. Use the sup to do theother tests required such as western, immunofluorescence and others. (If sup is
not used under sterile conditions, add 0.02% azide and keep at 4°C for months.)
6) Transfer the cells to a 25 cm2 flask in 4 ml of IMDM-m and continue to expand as
desired. The cells at this point are "addicted" to the macrophage supplement. It is
possible to wean them by gradually reducing the amount of MCM. We do it only ifvery large volumes are needed.
7) After deciding which one are the best clones, it is advisable to reclone them toascertain their homogeneity (next chapter).
8) The subclones should be checked by all the assays which were used to test theoriginals . The subclones should be frozen at least twice.
9) If convinced that the subclone is monoclonal, checking the isotype is possible . (Wehave been using a kit from Zymed - MONOAB-ID).
10) If not convinced that the hybrydoma is monoclonal, reclone a second time.

Cloning of Hybridoma
1) Make an accurate cell count of cells in log phase (3 X 105 - 1 X 106 cells / ml).
2) Determine the volume needed which will give you 100 cells.
3) Add the 100 cells to 25 ml of IMDM-m and plate 250 ml of the suspension in eachwell. Therefore, each well should get 1 cell delivered into it.
4) Clones should be visible and ready to screen in 10 to 14 days.

Ascites Production
1) 7 days before injection of cells, inject 0.5ml of Pristane (Tetramethyl - pentadecane,
Sigma T-7640), intraperitoneally.
2) Prepare cells in log phase.
3) On day of injection, spin 1-2 x106 cells per mouse.
4) Resuspend in IMDM or PBS at 1-2 x106 cells/0.5ml (do not leave the cells without
serum for more than 1 - 2hr). Transfer cells to syringe using a 16G needle.
5) Inject 0.5ml per mouse, using a 20G needle, intraperitonealy.
6) Collect ascites when animals are ready (swollen abdomen), 10 days to 15 days after
injection.
7) Spin fluid at 3000g for 10 min. to remove the cells. If there is an oil layer, remove it
and discard. Carefully remove the supernatant from the cells.
8) For storage, add sodium azide to 0.02%. Store at -20°C.

Literature
1. Antibodies - A Laboratory Manual; D. Lane, Ed Harlow.
2. Monoclonal Antibodies: Principles and Practice; James W. Goding.
3. Monoclonal antibody technology; Ailsa M. Campbell.

121

Polyclonal and monoclonal antibodies

122David Bowtell

ALKALI PLASMID PREPS

Large scale prep:1) Spin down cells and resuspend in 5ml (for a 50-200ml culture) of 25mM Tris, 10mM
EDTA pH 8.0.
2) Add 10ml of freshly prepared 0.2M NaOH, 1% SDS and mix thoroughly. Leave for 23min. MAXIMUM.
3) Add 7.5ml KoAc solution. Vortex.
4) Spin 10 min. at 7K in Sorval.
5) Pour supernatant through a piece of cheese cloth and add 11ml isopropanol. Spin 10
min. 7K in Sorval (the large amount of RNA acts as a carrier).
6) Resuspend pellet in 3ml of TE. Add to tube and make up to 4.3ml with TE. Add
100µl of 10mg/ml ethidium bromide and 4.4g of CsCl.
7) Band for 6hr (minimum) to o/n in VTi65 at 50-55K 20°C.
8) +/- reband by pulling band and adding to fresh tube and topping up with CsCl
density 1.6 (is 1g CsCl per ml TE, no extra E'Br required). Spin as above.
9) Remove ethidium bromide by extracting with saturated butanol 3-4 times. Isolate
plasmid DNA by adding an equal volume of TE and 5 volumes of room temperature
ethanol. Spin down plasmid immediately and resuspend in 400µl TE. Reprecipitate
in an ependorf tube by adding 1ml ethanol and 35µl 5M NaCl.

123

Alkali plasmid preps

Mini preps ("1 tube method")1) Grow overnight of 1.5ml.
2) Pour into eppendorf tube and spin down cells at 7-8K for 2min (low speed makes
them easier to resuspend).
3)Aspirate supernatant and resuspend cells in 50µl 25mM Tris pH 8.0, 10mM EDTA.
You can leave the lids off for all the following steps till you add the phenol.
4) Add 100µl of (freshly prepared) 1% SDS, 0.2M NaOH. Add it forcefully and you
don't need to vortex. It is not necessary to pause between steps 4, 5, 6.
5) Add 75µl KOAc solution and vortex.
6) Add 100µl of phenol/CHCl3. Close lids and vortex.
7) Spin in ependorf 13K for 2min.
8) Add supernatant to 500µl ethanol and spin 13K for 5min.
9) Aspirate supernatant, removing all the ethanol. Resuspend in 50µl TE (you don't
need a 70% rinse unless you are going to cut with a very salt sensitive enzyme).
10) Dispose of phenol into appropriate waste before discarding tubes.
I normally digest 2-5µl of this prep for an analytical gel. This DNA contains very little
chromosomal and is fine for cloning.
KOAc solution is:
60ml 5M potassium acetate
11.5ml glacial acetic acid
28.5ml of DDW.
Not necessary to pH it (it should be around pH 6).
No phenol method according to Gilles Morelle in Focus (11.1). At step 5 of mini-prep.
Add 75µl 7.5M ammonium acetate instead of KOAc (pH 7.8 without adjustment) andthen leave the tube on ice for 10 min.. The RNA and chromosomal will precipitate.
Spin 3min 10K and add 0.6vol. (100-125µl) isopropanol, leave on ice 10 min. and spin
15K 10 min., 70% rinse and resuspend pellet in 100µl TE.

124

Dan Kalderon

SMALL-SCALE PLASMID DNA PREPARATION
The fastest method is the "boiling method" but alkaline lysis (see large-scale
preparation of plasmid DNA) using 100µl sol'n I, 200µl sol'n II and 150µl sol'n III also
works well.
Boiling Method
Works for plasmid and M13 RF DNA
1. Grow saturated culture of plasmid-containing bacteria in 1.5ml L-broth or 2XYT
containing appropriate antibiotic with good aeration at 37°C or inoculate a 1:201:50 dilution of TG1(or DG98) O/N with a tooth-picked M13 plaque and grow for
5 hr.
2. Transfer most of the bacterial culture to a microfuge tube and pellet bacteria(30s).
Aspirate supernatant (store supernatant as a stock for M13 phage).
3. Resuspend bacteria in 100µl STET by pipette or vortex
STET is:8% sucrose
0.5% Triton-X100
50mM EDTA pH 8.0
10mM Tris-Cl pH 8.0
4. Add 100µl lysozyme (2mg/ml) in STET to each tube, mix and leave at room temp.
for 1-5 min.
5. Place tubes in boiling water bath (switch off heat to prevent tops popping) for 1min.
6. Spin 10 min at room temp. in microfuge.
7. Remove pellet with a flat toothpick.If the pellet occupies a large proportion of thevolume of the solution continue but add STET to 200µl and, next time, use morelysozyme; if the pellet is very small and brittle, use less lysozyme.
8. Add 30µl 4M ammonium acetate, 250µl isopropanol, mix and spin 10 min at room
temp in microfuge. Wash pellet with 70% ethanol, dry and dissolve in 20-50µl TE.
Enough DNA is recovered for at least 5 digests but the bulk of nucleic acid is RNA
which should be removed with DNase free RNase (made by boiling RNAase A at
2mg/ml for 30 mins. and cooling slowly) to see small DNA fragments. M13 R.F.
preparations usually contain a number of bands of lower molecular weight than
expected that presumably represent deletion products.

125Small-scale plasmid DNA preparation

9. Plasmid DNA prepared in this way can be used for
(i)
(ii)
(iii)
(iv)
(v)
(vi)

digestion with most enzymes
filling-in sticky ends with Klenow
purification of DNA fragments for ligation or for use as probes
nick-translation (with 32P or biotin)
double-stranded sequencing
template for riboprobe synthesis

Plasmid DNA prepared as described cannot easily be succesfully used for(i)digestion with several restriction enzymes, although digestion is normallyfine if the DNA is phenol(CHCl3/IAA) extracted and ethanol precipitated(although some members of the lab find this to be untrue).
(ii) Bal 31 digestion (except with good RNase treatment as RNA inhibits Bal 31
activity on DNA)(iii) Drosophila embryo injection(iv) in vitro transcription
Alternative methods from step 3:
A)

Add 350µl STET containing 1mg/ml lysozyme, resuspend thoroughly (1tube at a time), boil 2 min, spin, pick out snot(sic), add amm. acetate (this is
optional) and 350µl isopropanol, spin, wash in 70& ETOH and use 1/20 in adigest.

B)

As in A, but use 400µl isopropanol, skip the 70% ETOH wash, and use 1/50
per digest.

126

Ulrike Heberlein

MAXI PLASMID PREPS
For 500 ml cultures:1)

Grow 500ml cells at 37°C until OD600 rm = 0.6. Add 1.65ml of 50mg/ml
chloramphenicol in 100% ethanol to give a final concentration of 167µg/ml.
Incubate overnight (12-15 hours) at 37°C.

2)Harvest the cells at 5K for 10 min. Drain well.

3)

Suspend the pellets in 18ml of Solution I. Add 2 ml of Solution I containing
20mg/ml lysozyme (fresh). Incubate at room temperature for 10 min. (Note: if thecell culture was not amplified with chloramphenicol, use 40mg/ml lysozyme.)4)

Add 40ml of Solution II. Mix gently by swirling. Place in an ice water bath for 5
min.

5)

Add 20ml of pre-chilled Solution III. Mix the slurry by swirling. Incubate on ice for
15-20 min.

6)

Centrifuge at 10K in a GSA rotor for 10 min. Carefully decant the supernatant fromthe pelleted cell debris. Filter through a Kimwipe in a funnel to remove any debris.

7)

Add 0.6 volume of isopropanol (43.5ml). Mix by swirling. Let stand on ice for 20
min.

8)

Centrifuge for 15 min. at 10K in a GSA. Discard supernatant. Let the pellets dry
briefly (until the isopropanol has evaporated).

9)

Dissolve the pellet in 9.3ml TE. Add 10g CsCl and 0.7ml EtBr (10mg/ml).

10) Centrifuge at 38-40K, 20°C for 48 hr in 70.1Ti rotor.
11) Remove the bands (visualized by either long UV light or visible light) containing
plasmid DNA. Extract repeatedly (minimum 6X) with isopropanol (saturated with
CsCl and H2O) until the orange-red color of ethidium bromide is no longer visible.
12) Remove the CsCl by exhaustive dialysis overnight against TE at 4°C.
13) Add 1/10 volume of 3M NaOAc and 3 volumes of 100% ethanol. Chill at -70°C for
10 min and centrifuge at 17,000 x g for 15 min. Decant supernatant. Drain well.

127

Maxi plasmid preps

Solution I (50mM glucose, 25mM Tris pH8 10mM EDTA pH8)9.01g Glucose
25ml 1M Tris pH8.0
40ml 250mMEDTA pH8.0
DD H2O to 1 liter
Autoclave for 20 min.
Solution II
0.2M NaOH
1% SDS
Make fresh.
Solution III
600ml 5M Potassium Acetate (pH4.8) (= 300g/600ml)
115ml Glacial Acetic Acid
285ml DD H2O
Autoclave for 20 min.

128

Steve Mount

GROWTH OF M13 PHAGE
Growth and plaque-purification of M13 phage is generally performed as describedin the MRC protocols (Bankier and Barrell-1983-"Shotgun DNA Sequencing" inTechniques in the Life Sciences, B5; Nucleic Acid Biochemistry, B508, 1-34; Elsevier
Scientific Publishers Ireland Ltd.). The host strain is JM101 or TG1, which is isogenic
with JM101 except that hsd∆5 has been crossed onto it, rendering it Eco K r- m-.
Selection for retention of the F factor by growth on minimal media is essential for
JM101 and TG1. One should always grow fresh overnights in 2xYT from a colony on aminimal plate.
For preparation of RF, I quote: "When growing a bulk culture of M13 infected TG1,
10 ml. of a 1/100 dilution of an overnight culture is grown with shaking at 37° for 4 h.
[toothpick a plaque into 1.5 ml. of a 100-fold dilution of an O/N culture of TG1 in
2xYT broth; shake standing vertically in a rack at greater than 300 rpm until it isvisibly turbid (about 2 - 3 hours). Pour this infected culture into a 50 ml. tube holding
10 ml. of a growing culture of TG1, set up in parallel with the infected culture. Shakefor a further 4-5 hours.] After centrifuging for 10 min. at 2000 rev./min, thesupernatant containing the phage particles can be stored at 4° overnight [ We havefound that this sort of culture supernatant can be stored frozen at -20° following
growth in 2xYT for at least two years without significant loss of plaque-forming-units]
and used to infect a fresh culture of cells at A660 of ~ 0.2. This is grown with shaking
at 37° to A660 ~ 0.6 before proceeding with the chosen DNA purification technique."We've had good results allowing two 500ml. cultures to grow to A600 ~ 0.5 before
inoculating at 5 ml. per flask, allowing growth with vigorous shaking for another 4hours, then adding chloramphenicol at 25 µg/ml for the final 30 min. of growth.
Plaques are generally obtained by transfection with phage DNA. Dilute an
overnight 1/100 into 2xYT and grow for 1.5-2 h with shaking at 37° to A660 ~0.3.
Pellet cells at 4° and resuspend in one-half volume cold 50 mM CaCl2. Keep on ice 1530'. Spin again and resuspend in one-tenth of the original volume cold 50 mM CaCl2.
These competent cells do not freeze well, give the best results between 1 and 12 hours
after being made, and are not useful after 48 h. To transfect 200 µl of these cells are
aliquoted out into 13x100mm glass test tubes on ice, and ligation reactions or DNA (1µl of a 1/100 dilution of a standard single-stranded M13 miniprep works well) is
added directly, folowed by a 25-30 min. incubation. During this time top agar (0.7% in
1xYT) should be melted (at least 3 ml/plate). After the incubation the cells are heatshocked by placing the tubes in a 42° temp-block for 5 minutes. During the heat-shock
3 ml. top agar (50° C), 25 µl of the TG1 O/N, 25 µl BCIG ("X-Gal", 25 mg/ml in
dimethylformamide) and 25 µl IPTG (25 mg/ml filter-sterilized in water) per plate are
combined at 50° [of course, the indicator and inducer are only needed when screening
for inserts]; 3 ml. of this mixture is added to each tube, and the tubes are quickly
mixed and poured. Generally we use standard lambda plates, but plates of 1.2% agar
in 1xYT may give slightly better growth.
Single-stranded template DNA is prepared from plaques toothpicked into 2 ml.
aliquots of a 1/100 dilution of a fresh overnight placed in 13 x100 mm glass tubes.
These (multiples of 24 are most convenient) are grown standing in a vertical rack,129

Growth of M13 phage

shaking at greater than 300 rpm, for 4.5-6 h. at 37°. Transfer 1.5 ml. into a microfuge
tube and spin 5'. Carefully pour or pipet the supernatant into a fresh microfuge tube
containing 200 µl of 20% PEG, 2.0 M NaCl (polyethylene glycol 8000 from Sigma; thissolution should be filtered), being very careful not to carry over any of the pellet. Mix,and allow this to stand for at least 10 minutes. Spin 10 minutes in a microfuge.
Aspirate the supernatant, spin again for ~ 1 min, aspirate again. Resuspend in 100 µl
TE (be sure the pellet is suspended before adding phenol). The most rigorous
extraction procedure is to vortex well, spin for 5 min, and transfer the supernatant,using 1) 50 µl chloroform 2) 50 µl buffer-saturated phenol 3) 50µl of a 1:1
chloroform:phenol mixture. The simplest procedure is to use only phenol, and manychoose to do only the first two extractions. Following extraction, the aqueous phase isbrought to 150 mM NaOAc and precipitated with 250 µl of ethanol (usually overnightat -20°). The pellet is washed once with 95% ethanol and resuspended in 30-40 µl of
TE. This volume can be adjusted somewhat in response to the size of the PEG pellets.

130

Mark Fortini

GEL PURIFICATION OF OLIGONUCLEOTIDES
I generally purify 25-mer and larger oligonucleotides using the following
protocol. For smaller oligonucleotides, I use the TLC purification protocol (seefollwoing protocol). For many purposes, such as making primers for DNA sequencing,oligonucleotides may be purified sufficiently by passage through a G-10 spin column
after lyophilization.
1)

For gel purification, first dry down unpurified oligonucleotide solution in a
SpeedVac or lyophilizer.

2)

Redissolve the pellet in 50-100 µl 0.1 N NaOH. Add 1/3 volume of loading buffer(formamide/bromophenol blue dye).

3)

Prepare a 12% polyacrylamide gel (20:1 acrylamide:bis) as follows:-for 250 ml of gel solution (suitable for a 30 cm x 40 cm x 0.75 mm gel)
115 g urea
12.5 ml 20X TBE
63 ml ddH2O
75 ml 40% acrylamide solution (20 acryl:1 bis)
750 µl 25% ammonium persulfate
150 µl TEMED (add just prior to pouring gel)

4)

Cast gel using spacers of 0.75 mm thickness and wide-tooth combs (wells should be
2-4 cm wide).

5)

Load all or part of each oligonucleotide sample (from 2 above) onto the gel. Due todiffusion of the sample during electrophoresis, it is advisable to leave an empty lane
between each sample. Run gel at 30-50 watts until the bromophenol blue dye frontis ~75% down the gel.

6)

Separate the gel plates and transfer the gel onto Saran wrap. The oligonucleotide
samples are visualized by short-wave UV shadowing onto a TLC plate placed
under the Saran wrap and gel. The area of each gel lane containing theoligonucleotide appears as a large dark spot against the green illuminatedbackground of the TLC plate.

7)

Cut out the region of the gel containing the oligonucleotide with a razor blade.

8)

To elute the oligonucleotide out of the gel slices, cut each slice into ~1 cm2 pieces
and place all the pieces from each slice (usually 4-6) into 5 ml of elution buffer:
Elution buffer:
500 mM NH4OAc
1 mM EDTA
10 mM MgOAc
0.1% SDS

9)

Elute oligonucleotide out of gel for 4-6 hrs at room temp (gentle agitation is
optional). To check that elution is complete, examine gel pieces by short-wave UVshadowing as described above.

131Gel purification of oligonucleotides10) Extract the oligonucleotide eluate with phenol and ethanol precipitate. Wash thepellet 2X with 70% ethanol, dry and redissolve in a small volume (eg. 100-200 µl) ofTE, ddH2O or other solution.
11) Measure the concentration of the purified oligonucleotide solution using aspectrophotometer. For oligonucleotide solutions, I assume that an OD260 of 1.0 is
equivalentto40µg/ml.

132

Gel purification of oligonucleotides

Charles Zuker

PURIFICATION OF OLIGONUCLEOTIDES BY THIN LAYER
CHROMATOGRAPHY (TLC)
For practical purposes TLC-purified oligonucleotides are as good as gel-of-HPLCpurified oligos. TLC, however, is much cheaper.
1)

Lyophilize oligonucleotide (100-300µg); resuspend in 100-300µl of distilled water.

2)

Load oligo on a Silica Gel TLC plate (Silica Gel 60 F-254, EM Reagents, Cat. 5765-7,
20x20 cm) as a thin line across the plate approximately 2 cm from the bottom.

3)

Insert plate on a chromatography tank (30x30 cm) containing 100 ml running buffer(55ml n-propanol, 35ml NH4OH, 110ml H2O). Run until front is at or near the top.

4)

Remove plate and let dry inside a hood for a few hours (or until completely dry).
Visualize oligos using short-wave UV lamp and use a razor blade to scrape off youroligonucleotide (usually the lowest and most intense band of the ladder of oligos).

5)

Add silica powder to eppendorf tube containing 300-600µl of water, vortex for a few
min and spin down for 10 min. Re-extract the silica with 1/3 volume, spin, mix
supenatants and respin for 5 min.

6)

Read O.D. of your purified oligo (260 nm).

133

(D. Coit, P. Barr, P. Valenzuela)F. Laski
PROCEDURE FOR SITE-DIRECTED MUTAGENESIS
The following procedure is a modification of the primer-directed mutagenesis
technique developed in the Smith, Itakura and RajBhandary labs. It has been foundvery useful and is repeated here intact. Following the procedure are comments andmodifications which may be useful.
1)

Kinase the synthetic primer as follows: Mix 25 µl of primer in H2O (0.1 O.D.260
units, ~700 picomoles), 3 µl of 10X kinase buffer (500 mM Tris-HCl pH 8.0, 100mM MgCl2, 100 mM DTT), 1 µl of 100 mM ATP and 1 µl of T4 polynucleotide
kinase. Incubate at 37° for 30 min. Stop reaction by heating for 10 min. at 65°C.

2)

For annealing, mix 3 µl (2-3 µg, ~1 picomole) of M13 template DNA (or Bluescript
ssDNA - see comments below) containing the sequences to be mutated, 3 µl of
kinased primers (~70 picomoles) (directly from the above solution), 1 µl of asolution containing 200 mM Tris-HCl pH 7.5, 100 mM MgCl2, 500 mM NaCl and
10 mM DTT and 3 µl of H2O to complete 10 µl final volume. Heat at 55° for 5 min,then at 23° for 5 min.

3)

For extension and ligation, mix: 10 µl of the above annealing reaction, 10 µl of asolution containing 20 mM Tris-HCl pH 7.5, 10 mM MgCl2, 10 mM DTT, 1 mM
each deoxynucleotides and 0.1 mM ATP, 1 µl of a 1:2 dilution of BRL large
fragment E. coli DNA polymerase (10 units/µl) and 1 µl of Biolab's T4 DNA ligase(400 cohesive end ligation units/µl), in a total volume of 22 µl. Incubate at 16° for
18-24 hrs.

4)

OPTIONAL: For treatment with nuclease S1 mix: 2 µl of the solution after
extension and ligation, 1 µl of 10X S1 buffer, 1 µl of a 1:100 dilution of BRL S1nuclease diluted in 1X S1 buffer, and 6 µl of H2O to complete 10 µl final volume.
Incubate at 37° for 15 minutes.

5)

For transformation, mix: 10 µl of the S1 reaction mix and 100 µl of JM-101
competent cells. Let sit on ice for 60 min. For plating, add 200 µl of log phase
JM101 in YT medium and 3 ml of 0.8% agarose in YT, preheated to 45°. Mix andplate over preheated (37°C) agar plates. Incubate at 37° overnight.

6)

For screening by plaque hybridization: Cool plates in the cold for 60-90 min.
Overlay a nitrocellulose filter over the plaques, press slightly and leave for 10 min.
Carefully lift filters away from plate and leave them drying at room temperaturefor 60 min.

7)

For lysis and fixing place filters over 3 M filter paper moisted with the following
solutions: 5 min over 0.5 N NaOH-1.5 M NaCl, 5 min over 3 M NaCl-1M Tris-HCl(pH 7.5), 5 min again over 3 M NaCl-1 M Tris-HCl (pH 7.5) and 5 min over 2X
SSC. Bake filters for 2 hrs at 80° under vacuum.

8)

For pre-hybridization, incubate each filter in 5 ml of 0.2% SDS, 10% Denhardt's
solution, 6X SSC at room temperature for 60 min.

134

Procedure for site-directed mutagenesis

9)

For hybridization wash filters in 6X SSC for 5 min and incubate each filter in 5 mlof 10% Denhardt's, 6X SSC and 2x105 cpm of probe fragment (probe DNA
fragment is kinased with γ-ATP to specific activity of 1-4x107 cpm/µg). Incubate
at room temperature for 2 hrs.

10) Wash filters in 6X SSC at room temperature for 5 min. Expose to X-ray film for 2
hr with screen. To differentiate specific from non-specific hybridization wash
again at higher temperatures, i.e. 40°, 55° etc. (depending on the length andcomposition of the probe), and expose to X-ray film.
11) For plaque purification, take the agar region containing the positive plaque andsuspend in 5 ml of YT. This should contain a total of 108 to 109 pfu. Dilute thissolution 200 times in YT to make a stock of approximately 106 pfu/ml. Dilute thisagain 100 times in YT to approximately 105 pfu/ml. Plate 10, 50 and 100 µl as
described in step 5.
12) Repeat steps 6 to 10 as necessary.
13) Take isolated plaque and suspend in 1.5 ml of a 1:40 dilution of log phase JM101
in YT. Grow for 5 hrs at 37°. Centrifuge in 1.5 ml Eppendorf tubes. Savesupernatant for DNA sequencing and pellet for alkaline-SDS miniscreens.
Comments and Changes on Procedure
Purification of M13 and Bluescript Single Stranded DNA
This is my procedure for preparing M13 single strand DNA, but feel free to use
your normal procedure. One ml of a TG1 overnight is added to 300 ml of 2xYT.
Thirty minutes later add 20 ul of your M13 recombinant phage. Shake for ~ 7 hours,
then spin down bacteria for 10' at 6,000 RPM. Add to supernatent (300 ml) 20 ml of 5
M NaCl and 10g PEG. After PEG goes into solution let it sit for 10 min. on ice. Spin 10
min. at 6,000 RPM, pour off supernatent and let the pellet air dry. Resuspend thepellet in ~5 ml of TE, phenol 2 times, chloroform 1 time and ethanol precipitate.
Resuspend the pellet in TE (as small a volume as easily possible, usually between 400
and 800 µl).
Many people use this M13-recombinant single strand DNA directly formutagenesisis. I however, first clean it up over an alkaline sucrose gradient. The
above single stranded DNA is contaminated with short fragments of DNA and/or
RNA that can prime the single stranded DNA in the Klenow reaction, resulting inincreased background and possibly double mutations. These short fragments are
removed on the alkaline sucrose gradient, which results in DNA that is primed only if
an exogenous oligonucleotide is added.
The alkaline sucrose gradient procedure is as follows: one half ml of a solutionthat is 50% sucrose, 1 M NaCl, 0.2 N NaOH 1mM EDTA is added to a 1/2 inch
diameter x 2 inch polyallomer tube. A 5% to 20% gradient is then poured ontop of the
50% sucrose solution. This is done by putting 2.3 ml of a 20% sucrose solution (that is
1M NaCl, 0.2 N NaOH, 1 mM EDTA) into one slot of a gradient former (the slot closest
to the spout) and 2.3 ml of a 5% sucrose solution (1 M NaCl, 0.2 N NaOH, 1 mM
EDTA) into the other. The gradient is then poured, don't worry if it isn't perfect. 50 to200 µl of the single stranded DNA solution (100 to 200 µg) in TE is pipetted onto the
top of the gradient. The tubes are then spun for 5 hours at 45,000 RPM at 16° C in a
pre-cooled SW 50.1. The gradient is then dripped from the bottom in 200 to 250 µl

135Procedure for site-directed mutagenesis

fractions and the fractions assayed by O.D. The peak 2 to 4 fractions are usually at
fraction 9 or 10. The purified DNA is neutralized by adding 1/10 volume of 1M tris
pH 8.0 and 1/10 volume of 2 M HCl, and thenn 2.5 volumes of ethanol added and the
DNA precipitated. The DNA is resuspended in TE and brought to a concentration of 1
mg/ml. Final recovery is usually between 30-50% of input DNA.
This mutagenesis protocol also works well for single-stranded templates in theBluescript vector system (Stratagene, Inc.). Single-stranded Bluescript DNA isprepared as for DNA sequencing according to the Bluescript protocol booklet. The
yield from a small-scale ssDNA prep (1.5 ml culture) suffices for 2-3 mutagenesis
reactions.
Comment on Kinasing Oligonucleotide
My procedure is as follows: The oligonucleotide is diluted into the 30 µl
reaction so that the O.D. 260 of the oligonucleotide in the kinase reaction is 4.0. As an
example, if the O.D. 260 of a 1/100 dilution of the oligonucleotide is 0.295, the the O.D.
260 of the oligonuleotide is 29.5 and I want to dilute it by a factor of 0.135 (4.0/29.5)
and therefore will add 4.05 µl (0.135 x 30µl) to the 30 µl reaction. I will of course roundthe 4.05 µl to 4 µl. Everything is then done as above except that I only do one half hour
reaction and I don't bother to kill the enzyme at 65°C.
Comment on Annealing
I do exactly as they suggest except that I heat the sample at 65° instead of 55° (Idon't have a 55° water bath) and I allow hybridization at room temperature instead of
23° (I don't have a 23° water bath).
Comment on Extension and Ligation
I add 1/2 µl of both Klenow and ligase without diluting it.
Comment on S1 Treatment
Instead of just treating all the DNA with S1 nuclease for 15 min., I always
perform a titration, removing 2.5 µl aliquots of the reaction at 0, 1, 5 and 15 min. I findthat for some reactions, a 15' incubation is too long and you get no plaques. The
titration allows you to knock down the background but still retain enough plaques towork with. I should also add that you can skip the S1 treatment altogether, although
this will usually decrease the number percentage of phage carrying the mutation.
However, when using an oligonucleotide that will create an insertion or deletion I
sometimes find that S1 treatment will decrease the percentage of plaques that containthe desired mutation, presumably because the S1 directly digests the extended andligated DNA at the loopout caused by the oligonucleotide.
Comment on Transformation and Plating
I follow my normal transformation procedure: mix 2.5 µl of the S1 reaction mix
and 100 µl of TG1 competent cells. Let sit in ice for 10 min. Heat at 37° for 5 minutes.
Add 1 ml of 2xYT at room temp. For plating, mix 200 µl of the M13-transformed TG1
cells with 100 µl of TG1 overnight and 3 ml of 0.8% agarose in YT, preheated to 45°.
Mix and plate over agar plates. Incubate at 37° overnight. Once the plaques arevisible, plate out all or some of the remaining M13-transformed TG1 cells (which havebeen stored at 4°C) so that the plaques will be well separated. I find that between 40 to~150 plaques per plate are ideal.
For Bluescript templates, you should do a standard bacterial transformation of
the S1 reaction mix and plate out on Amp plates to obtain colonies rather than plaques.

136Procedure for site-directed mutagenesis

Properly mutagenized templates are then detected by colony hybridizations rather
than plaque hybridizations, using the same conditions as described in steps 6-10
above.
Comment on Making Probe
I mix 1 µl of oligonucleotide (diluted to O.D. 260 = 1) with 1 µl of 10x kinase, 2 µl
32
of
P-ATP (10mCi in ~50µl, carrier free from ICN), 6µl of water and 1/2 µl of kinase.
Incubate for 30 min. at 37°C. I use 1/2 of this mixture directly for screening, you don't
have to put it over a column.
Comment on Plaque Purification
Even if it is obvious which plaque contains the transformant I always plaquepurify it because some of the plaques contain a mixture of mutated and wild typegenomes. Instead of diluting to plaque purify I streak out the plaque in the samemanner you would a bacterial colony and then very gently add 3.5 ml of top agar
containing 100 µl of a TG1 overnight.
Comment on RepeatingThis is usually not needed.
Comment on Sequencing
I usually prep 4 plaques for sequencing.

137Procedure for site-directed mutagenesis

from Grace Gill via Betsy O'Neill

SITE-DIRECTED MUTAGENESISUse of the strain CJ236 (ung- dut-)to make the single stranded template DNA sitedirected mutagenesis causes dUTP to be incorporated in place of dTTP. Later in theprocedure after the mutation containing primer is annealed, the second strand issynthesized in vitro and the plasmid is transformed into TG-2 cells, the repair
mechanisms in the cell tend to repair the uracil-containing parental strand, thusincreasing the frequency of appearance of the desired mutant. Reports indicate anincrease from about 5% to about 70% (Biotechniques vol. 5, pp786-791, 1987).
1) Purify oligo by TLC or electrophoresis.
2) Kinase 50 pmole of oligo in a 25µl reaction.
3) Set up 2 annealing reactions, one with primer and one without primer.
ssDNA

1µl (see other protocol)

primer 1 or 0 µl (2pm kinased oligo)
10x anneal
1 µl
ddH2O to 10 µl
4) Heat to 70°C/10 minutes. Cool slowly to 30°C, then place on ice.
5)

For extension reaction, add to primer annealed template:
10x extens 2µl
dNTPs 4µl

(2.5mM each)ligase 1µl(dilute concentrated NEB stuff 4x)

T4 Pol 1µl
ddH2O

2µl

6) Leave on ice 5 minutes. Transfer to RT 5 minutes. Incubate at 30°C/90 minutes.
Lower temperatures may be necessary for primers with a lot of mismatches.
7) Bring volume to 100ul with TE. Phenol/cholroform extract, ethanol precipitate.
8) Transform 1/10th of reaction into TG2.
10x Annealing Buffer

10x Extension Buffer

200 mM Tris, pH 7.5
20 mM MgCl2

10mM ATP100 mM Tris, pH7.5

500 mM NaCl

50 mM MgCl2
20 mM DTT

138

Procedure for site-directed mutagenesis

Ulrike Heberlein

DOUBLE STRANDED SEQUENCING
Minilysate DNA:
1)

Prepare DNA using either alkaline lysis or STET/boiling protocols from 1.5 ml ofsaturated bacterial culture. (I have found that DNA isolated from strains HB101 or
TG-2 sequences well, whereas DNA isolated from TG-1 doesn't work as well).
Never freeze the bacterial pellet before isolating the DNA!!! Before denaturing,clean the DNA by consecutive phenol:chloroform and chloroform extractions.
Precipitate with 2.5 volumes of ethanol in the presence of 0.3M NaOAc and wash
DNA pellet with 70% ethanol. Resuspend in 25µl of TE.

2)

To 5µl of the cleaned DNA add 0.5µl of 0.5mg/ml RNAse A and incubate at room
temperature for 2-5 minutes.

3)

Add 5µl of 1M NaOH (make up from 10M stock about once a month) and incubate
at room temperature for 5 minutes.

4)

Neutralize by adding 3µl of 3M NaOAc pH 5.0.

5)Add 7µl of a ~1pmole/µl solution of primer (MW is~330g/mole/base; absorbanceof a 1mg/ml single stranded DNA solution is ~25).

6)

Precipitate with 75µl of 100% ethanol at -70°C for 15-30minutes (or in dry ice for 510 minutes). Spin in a cold microfuge for 20 minutes. Wash the pellet with 70%
ethanol and dry (the pellet should be visible).

7)

Resuspend thoroughly in 8µl of water, add 2µl of Sequenase buffer and anneal as
usual. Follow Sequenase instructions.

CsCl purified DNA:• Use 2-3µg of DNA in a 5µl volume. Denature with 5µl of 1M NaOH for 5 minutes atroom temperature and follow protocol described for minilysate DNA.

139

Don Rio

PREPARATION AND PURIFICATION OF E. COLI β-GALACTOSIDASEFUSION PROTEINS AND THEIR USE AS IMMUNOGENSIntroduction
We use the pUR series of plasmid vectors (Ruther and Müller-Hill, EMBO J.
(1983)). These plasmids carry polylinkers in the C-terminal portion of the βgalactosidase gene beyond the Eco RI site used for insertion in lgt11. Therefore, thefusion proteins will generally retain β-galactosidase enzymatic activity. This could be
useful for following purification or for affinity chromatography. In any case, select theappropriate vector and clone your open reading frame segment (generally 200-500 bp)
into the polylinker using E. coli strain 71-18. Analyze the clone by restriction mapping
and/or DNA sequencing to confirm insert orientation and maintenance of the open
reading frame.
Analytical analysis of fusion proteins:
The idea here is to examine the size of your fusion protein and to determine its
abundance and stability.
1)

Grow 1 ml overnight culture in L-broth with 100 µg/ml ampicillin (no glucose).

2)

Inoculate 2 fresh 1 ml cultures with 50 µl of o/n. Grow until A590 ~0.2-0.4 at
37°C (~2 h).

3)

To one culture add IPTG to 1 mM (1:100 of 25 mg/ml). Incubate at 37° for 2 h.

4)Spin down cultures in 1.5 ml Eppendorf tube and drain liquid.

5)

Resuspend pellet in 100 µl 2X SDS gel sample buffer (use Eppendorf mixer andresuspend completely: 20-30 minutes).

6)

Boil 2-4 min in boiling H2O broth.

7)

Spin 5 min Epp centrifuge.

8)

Load 10 µl on 7.5% SDS polyacrylamide gel.

9)

Stain gel with Coomassie blue.

10)

Destain gel and visualize bands.

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Preparation and purification of E. coli β-galactosidase fusion proteins and their use as immunogens

Preparative fusion protein purification
Plan A: Preparative SDS-gel electrophoresis (this is suitable for unstable fusion
proteins).
1)
Grow 100 ml IPTG-induced culture, harvest by centrifugation and resuspend in
3-5 ml 2X SDS-gel sample buffer.
2)

Boil 5 min and spin at 12,000 x g for 10 min.

3)

Remove supernatant and load ~1 ml onto 1.5 mm thick 7.5% preparative SDS
gel.

4

Stain gel with 0.25 M cold KCl. The fusion protein band appears as a whiteband where K+-SDS complexes precipitate. Alternatively, the gel can be stainedwith Coomassie blue without acetic acid and then detained in H2O.

5)

Excise band and either pulverize it for direct injection or electroelute the protein
and concentrate it by acetone precipitation.

Plan B: Insoluble aggregate purification. (This is suitable for abundant fusion
proteins and when you
want a lot of fusion protein, e.g. for coupling to
CNBr-Sepharose for affinity purification).
1)

Grow 1 L IPTG-induced culture; harvest by centrifugation and freeze at -70°C.

2)

Thaw cell pellet on ice in 50 ml Buffer A (50 mM Tris-HCl pH 7.9, 200 mM NaCl,
2 mM EDTA, 2 mM b-mercaptoethanol, 100 µg/ml PMSF, 100 µg/ml PMSF, 1µM pepstatin and 1 µM leupeptin).

3)

Add solid lysozyme to 0.2 mg/ml - incubate on ice for 20 min.

4)

Add triton X-100 to 1% - incubate on ice for 10 min.

5)

Add Zwittergent 3-14 (CalBiochem) to 0.5% . Incubate on ice for 10 min.

6)

Sonicate 3 x 15 seconds on ice using a Branson sonicator with a microtip.

7)

Layer the lysate onto 10 ml of 40% sucrose in 10 mM Tris-HCl pH 7.5, 1 mM
EDTA, 0.2 M NaCl.

8)

Centrifuge 30 min at 13K rpm in a Sorvall HB-4 swinging bucket rotor.

9)

Resuspend aggregate pellet in 2 ml PBS.

10)

Add 20 ml urea extraction buffer (8M urea, 0.5 M NaCl, 0.5 M Tris-HCl, pH 7.9,
1 mM EDTA, 30 mM b-mercaptoethanol, 1 µM pepstatin and 1 µM leupeptin).

11)

Vortex vigorously until pellet is dissolved.

12)

Dialyze at room temperature for 2 h against 2 L of 50 mM Tris-HCl pH 7.9, 0.5
M NaCl, 10% glycerol, 200 µg/ml PMSF with 1 change.

13)

Remove insoluble material by centrifugation at 10,000 x g for 10 min.

141

Preparation and purification of E. coli β-galactosidase fusion proteins and their use as immunogens

14)

The fusion protein is present in the supernatant at ~1-4 mg/ml and is ~80%
pure. It can be used directly for immunization and should be stored at -20°C.

Immunization:
1)
We use 3-5 lb. female New Zealand white rabbits.
2)

Inject 50-200 µg fusion protein in 0.5 ml PBS + 0.5 ml complete Freund's
adjuvant (Gibco) at multiple intradermal sites or subcutaneously.

3)

At 3-week intervals - a boost of the same amount of antigen with incomplete
Freund's adjuvant. After 2 boosts, the serum is usually high titer. The rabbits
are bled and serum prepared.

4)

I have kept boosting for 3-4 months and can still get usable (high titer ) serum.

Preparation of Serum from Blood:
1)
Bleed rabbit from central ear artery into glass Corex tube. Allow blood to clot atroom temp for 1-2 h.
2)

Use pipette to dissociate clot from sides of glass tube and leave o/n at 4°C forclot to contract.

3)

Use pipette to remove serum (should be approx. 1/3 of the total blood volume).
Spin at 1000 rpm in benchtop clinical centrifuge or 5K in Sorvall SS34 for 5 min
to pellet RBC. Store at -20°C in conveniently sized aliquots.

142

Kevin JonesFUSION PROTEINS FOR ANTIBODY PRODUCTION
This method for the production of specific antisera includes purification of afusion protein produced in bacteria by isolation of an insoluble protein fraction (fusion
proteins expressed at high levels frequently form inclusion bodies which are
insoluble), further purification by preparative gel electrophoresis (SDS-Laemmli
method), electroelution of the protein, and injection into mice (or other appropriate
animal — the general experience of members of the Rubin lab has been that mice often
produce quality sera having low background). Mice are also a good choice if one
might be interested in generating a monoclonal.
With respect to the fusion protein vector selection, there are many possible
choices and the ideal choice may be quite dependent on your protein. The pATH
fusion vectors (trpE fusion) have worked quite well for me, I have obtained a mg ofpure protein from 50ml of culture with several different constructs using these vectors
and the method described below. Other reports of similar yields with trpE fusions arepresent in the literature. The pUR vector series (lacZ fusion) has been successfully
used by many people in the Rubin lab, the primary advantage of this and other lacZ
fusion vectors is that they lead to a very large fusion protein (116kd + what you add)
which is substantially larger than most of the other proteins which are present in abacterial extract, facilitating purification by gel electrophoresis. lacZ fusion proteinsare also often insoluble or if soluble can be purified with a substrate analog affinity
resin. Yields per ml of culture will generally be substantially lower than thoseobtainable with trpE fusions. Several lacZ fusion vector series other than the pUR
series are availible at this writing, and they may provide superior yields, but I have not
had any experience with these. The T7 vector system (pET vectors) deserves a mention— this vector system is useful for producing protein initiated at the bona-fide
initiation codon, or for making fusion proteins (to a portion of a T7 coat protein).
Levels of protein production comparable to that obtained with trpE fusions can beachieved. In summary, with regard to selecting vectors my suggestion is that you
choose two different vectors which have convenient restriction sites and which
produce protein having the desired characteristics (i.e. intact or fusion, methods ofpurification). It is useful to have fusions to different proteins (e.g. lacZ and trpE) so
that a sera raised against one fusion can be tested with the other, or one fusion protein
can be used for affinity purification of sera raised against another.
The choice of regions in your protein to use as antigen is somewhat arbitrary but
there are several points worth mentioning. First, hydrophilic regions are often morestrongly antigenic than are hydrophobic regions. Second, it is wise to try several
different (preferably non-overlapping) regions if possible. Yields of fusion can be quite
variable with different constructs, and one may also be a lot more antigenic thanothers. Also, if you have several sera directed against different portions of the protein
which give identical results in histochemical, western experiments, etc. it is likely yoursera are recognizing the protein you hope they are (in the absence of better controls).
The protocol below is based upon methods described for purifying insoluble
trpE and lacZ fusions. Therefore, while the conditions for the growth of cells andinduction are described for trpE fusions, it should in principle be applicable topurification of any insoluble protein fusion. For general information about production
and use of antibodies, the laboratory manual by Harlow and Lane is highly
recommended.

143

Fusion proteins for antibody production

FUSION PROTEIN PURIFICATION-trpE fusions
1) Pick colony, grow overnight in L-broth with 100mg/ml AMP.
2) Inoculate M9-Casamino Acids media (100mg/ml AMP) 1:100; grow to O.D.600 of
approximately 0.2 and add a 1:1000 dilution of 10mg/ml β-indoleacrylic acid
(Sigma). Let grow overnight (8-16hr).
3) Chill cells on ice, harvest 4krpm, 10'.
4) Store cell pellet at -80o.
(Volumes given in parentheses after instruction are for 25ml, 50ml, and 500ml cultures,
respectively.)5) Thaw cells on ice in 1/10 vol TEN (culture vol=1 vol), resuspend by vortexing.
(2.5ml, 5.0ml, 50ml)
6) Add lysozyme to 0.2mg/ml. (50µl, 100µl, 1ml)
7) Let sit on ice 20'.
8) Bring to 0.2% NP-40,10' on ice. (add 100µl, 200µl, 2ml)9) Add .15 x culture vol 1.5M NaCl, 12mM MgCl2. (3.75ml, 7.5ml, 75ml)
10) Add DNase I to 4 µg/ml, leave on ice 1hr, stir regularly.
11) Layer onto sucrose cushion (at least 1/100 culture vol). Spin 30' 13krpm in Sorvall
HB-4 (swinging bucket) or equivalent.
12) Resuspend aggregate pellet in TEN and re-spin (to wash). This wash can be
repeated several times if desired.
13) The protein can be run over a prep gel; at the end of step 12 it should be at least
50% pure.
14) An extra step number, for luck.
Solutions:
TEN: 0.2 M NaCl, 10 mM Tris pH 8.0, 1 mM EDTA
sucrose cushion: 40% sucrose in TEN (filter sterilize and store at 4oC)
Assorted Notes:
1) Sonication can be substituted for DNase treatment (steps 9 and 10), both appeared
to work equally well in my hands. I found DNase treatment to be easier.
2) Protein can be electroeluted from the gel slice in protein gel running buffer and thendialyzed against 1xPBS. After quantitation, this material can be used for

144

Fusion proteins for antibody production

immunization. Generally, about 20mg/mouse is considered plenty for each
immunization.
3) I found the use of protease inhibitors to be unnecessary. You may need them foryour fusion, but since they are often nasty or expensive, I'd omit them if possible. It
would seem likely that protein in inclusion bodies might be somewhat resistant toproteolysis, so any degradation products you see (you probably will see some) may
be a result of degradation of protein during the period the cells are actively
synthesizing it, and not during your purification.
REFERENCES
Science 214:1125 (1981)
J. Virol. 49:132(1984)
Cell 44:21 (1986)
Antibodies-A Laboratory Manual; Harlow, E., Lane, D. (1988) Cold Spring Harbor
Laboratory Press

145

Kevin Moses

MAKING Trp E FUSION PROTEINS FOR ANTIBODY PRODUCTION
This procedure aims to make a novel protein with the N-terminal 37 kD of the E.
coli Trp E gene product and any other protein at the C-terminus in order to inocculate
animals for the production of anti-sera. In outline you must insert DNA bearing youropen reading frame into the polylinker of one of the pATH vectors such that yourprotein will be appended to the first 37 kD of the bacterial trpE gene product. You will
make a translational fusion which must therefore be in frame. There are several pATH
plasmids with various polylinkers (see map/sequences below) and at this time all areunpublished. Details may be obtained from one of their creators, Alex Tzagolof(Columbia (212) 854-2920). I have used the trio pATH 1, 10 and 11 which supply EcoR1
sites in each of the three frames:
pATH 11:
pATH 1:
pATH 10:GAA TTC CAGCGA ATT CGA
CCG AAT TCG

Anecdotally pATH11 gives the highest protein yields. You can choose appropriate
cDNAs that end at a 5' synthetic Eco R1 site, or engineer one in by site directedmutagenesis, or use a naturally occurring site. Otherwise there are many otherrestriction sites in the polylinkers to choose from. Don't worry about the C-terminus, as
all three frames stop soon after the polylinker. All of these vector DNAs grow very
poorly, so be prepared to put all of your miniprep plasmid DNA in one track to see it ona gel. Including tryptophan in the growth medium may help avoid selecting nonexpressing mutants. Once you have your subclone in E. coli HB101 do the following:
1) Grow a 5ml seed culture in LB + 50 µg/ml ampicillin shaken at 37°C to saturation(overnight).
2) Inoculate 250 ml of M9CA + 50 µg/ml ampicillin in a 2 liter flask with 2.5 ml of theseed culture and grow shaken at 37°C to A600 = 0.2 (about two hours).
3) Add 250 µl of 10 mg/ml 3β-indole-acrylic acid (Sigma No.I-1625) in propan-1-ol andcontinue to incubate shaken at 37°C for at least four hours (I usually go overnight).
4) Make up 50 ml of TEN+ and store on ice.
5) Spin down the cells and resuspend in 25 ml TEN+ and transfer to a 50 ml disposable
polypropylene tube, add a dash of lysozyme (Sigma No. L-6876) and incubate on ice
for 25 minutes.
6) Add 2.5 ml of a 10% solution of triton X-100 (Sigma No. T-6878) in water, vortex andincubate on ice for 10 minutes.
7) Add 1.25 ml of a 10% solution of Zwitergent (Calbiochem No. 693017) in water, vortex
and incubate on ice for 10 minutes.
8) Sonicate at maximum microtip energy in repeated 10 second bursts with cooling on
ice for a total of 60 seconds, or until the viscosity drops significantly. This step breaks

146

Making Trp E fusion proteins for antibody production

E.coli DNA, which otherwise reduces the effects of the subsequent wash steps andcan interfere with the running of preparative gels.
9) Keep 50 µl of the total extract and load the rest carefully (so as to avoid mixing) onto a
10 ml sucrose cushion (40% sucrose in TEN) in a 50 ml polycarbonate tube. Mark theposition of the interface.
10) Spin for 30 minutes at 4°C at 12,000 rpm in an HB-4 rotor (or similar).
11) Take off the supernatant above the interface, keep a 50 µl aliquot handy and store therest frozen at -20°C. Then remove and discard the sucrose.
12) Resuspend the pellet in 5 ml TEN+ and transfer to a 15 ml corex tube on ice. To breakup the pellet you will need to thoroughly beat up on it, vortex it, draw it through apipette tip or use any other violent means.
13) Spin at 9,000 rpm at 4°C for 5 minutes, and remove and discard the supernatant.
14) Repeat steps 12 and 13 until the supernatant becomes clear or you think you will runout of TEN+, or you get bored. I usually do three such washes.
15) Resuspend the pellet in 2 ml TEN+, keep a 50 µl aliquot handy and store the rest at 20°C.
16) Pour a 9% acrylamide SDS gel at pH 8.8 with a pH 6.8 stacker. For my size apparatus
these are as below, but can be scaled to fit:
9.6 ml 30:0.8 acrylamide:bis
12.0 ml 1 M Tris pH 8.8 (from base)10.0 ml distilled water
320 µl 10% SDS 100 µl 10% SDS
160 µl fresh 10% Ammonium persulfate
30 µl TEMED

1.66 ml 30:0.8 acrylamide:bis
1.25 ml 1 M Tris pH 6.8 (from base)
6.92 ml distilled water
50 µl fresh 10% Ammonium persulfate
12 µl TEMED

17) Load 15 µl each of the total and supernatant and 5 µl of the pellet in equal volumes of
2X Lämmli buffer. Also run appropriate size standards (such as Sigma No. SDS-6H).
It is a good idea to do a parallel prep on induced vector alone so that you can see the
37 kD TrpE protein.
18) Run the gel as fast as you can while keeping it cool in 1X PGB and then stain for one
hour in coomassie stain.
19) Destain repeatedly until satisfied with the result (overnight works well), and dry thegel down onto 3MM paper.
20) Comparison between the vector and insert preps should immediately reveal yourfusion protein when the gel is stained, hopefully at the correct molecular weight. If
you can't see anything quit now and get a steady job. If you have what you want,scale up the gel and run the pellet out on a prep gel.
21) Stain the prep gel for only two minutes in aqueous coomassie, and destain
extensively in water.

147

Making Trp E fusion proteins for antibody production

22) Put the gel in a pyrex dish and view it on a white light box. Cut out the band andelectroelute in your favorite apparatus in 1XPGB in a cold room. You can follow thepassage of the coomassie blue.
23) Dialyse extensively (at least four changes over 24 hours) against 1X PBS + 2mM
β−mercapto ethanol (140 µl per 1 liter) + 1.74 µg/ml PMSF.
24) Run an aliquot adjacent to a dilution series of a known protein (eg BSA) and assess
its concentration.
25) Store in aliquots at -20°C. This is your antigen.
Buffers (in order of use)
LB - see buffers/media section
M9CA - see buffers/media section
TEN+ (50 ml)
0.5 ml 1 M Tris pH 8.0
0.2 ml 250 mM EDTA
2.0 ml 5 M NaCl
7 µl β-mercapto ethanol
50 µl 1.74 mg/ml Phenylmethane-sulfonyl, in propanol (Boehringer Manheim)
50 µl 1 M Benzamidine in water (Sigma B-6506 stored at -20°C in 1 ml aliquots)
50 µl 2 mg/ml aprotinin in water (Boehringer Manheim stored at -20°C in 1 ml aliquots)
50 µl 1 mg/ml leupeptin in water (Boehringer Manheim stored at -20°C in 1 ml aliquots)
50 µl 1 mg/ml pepstatin in methanol (Boehringer Manheim stored at -20°C in 1 ml aliquots)
distilled water to 50 ml
10% triton X-100 (Sigma No. T-6878) in water
10% Zwitergent (Calbiochem No. 693017) in water148

Making Trp E fusion proteins for antibody production

40% sucrose (in TEN)
1.0 ml 1 M Tris pH 8.0
0.4 ml 250 mM EDTA
4.0 ml 5 M NaCl
40 g sucrose
distilled water to 100 ml, filter sterilise and store at 4°C.
2X Lämmli buffer
20 % glycerol
4 % SDS
250 mM Tris pH 6.8
10 % β-mercapto ethanol
0.05% bromophenol blue
5X PGB (Protein Gel Buffer)30.27 g Tris base144.1 g glysine
10
g SDS
water to 2 liters, store at room temperature
stain - 50 ml glacial acetic acid (to 10%)125 ml Ethanol (to 25%)
0.25 g coomassie brilliant blue (Sigma No. B-1131)
distilled water to 500 ml, store at room temperature (reusable)
destain -Same as stain less the coomassie

aqueous coomassie
0.25 g coomassie brilliant blue (Sigma No. B-1131)
distilled water to 500 ml, store at room temperature (reusable)
1X PBS
0.1 M sodium phosphate buffer pH 7.2
0.7 % NaCl

149

Zhichun Lai

GST FUSIONS FOR ANTIBODY PRODUCTION
pGEX is a protein expression vector for use in E.coli that carries the GST(glutathione S-transferase, 26 Kd) gene from the parasitic helminth Schistosoma
japonicum. A gene encoding a foreign protein is fused at the terminus of the GST gene ata polylinker (BamHI, SmaI, EcoRI). Each of the three vectors, pGEX-1, pGEX-2X andpGEX-3X, generates fusions in a different ORF. The GST gene is under control of the tac
promoter and is induced by IPTG. Plasmid derived lacq product is sufficent to inhibt
the GST gene expression in the absence of inducers.
Most GST fusion proteins are soluble, and one may purify these GST fusion
proteins by affinity to glutathione, the substrate of the enzyme, linked to agarose beads.
Supernatant containing GST or GST fusion protein is mixed with the beads. Afterextensive washing, the adsorbed protein is released by adding free glutathione ascompetitor. This one step purification gives pretty pure material suitable for injection
into an animal, or for biochemical analysis. GST fusion proteins are assumed to keep
their native comformation, since no denaturation steps are involved.
The yield is about 1.6-15 mg/liter culture.
Mini Prep:
1) Screen TG2 transformant.
2) Take 100 µl overnight culture into 900 µl 2xYT with 100 µg/ml ampicillin.
3) Grow at 37°C for 1.5 hours; add IPTG to final concentration of 1.0 mM; grow 1.5 hours
at 37°C. Spin down cells, resuspend in 300 µl PBT buffer (PBS + 1% Triton X-100)with 0.2 mg/ml lysozyme; sonicated about 5" (not more than 10"); spin down debris;mix supernatant with 50 µl 50% glutathione-agarose beads; leave at room
temperature for a few minutes; wash 3x 1ml PBT; add 30 µl 3x SDS sample buffer
and boil for 5 min. before loading to SDS protein gel.
Comments: If you don't see any fusion protein with expected size, you may run total
protein on the gel. Use the one without IPTG as a control. It is possible that fusion
protein is insoluble. If you still don't see anything, just simply give up, try otherconstructs, or other expression systems.
Maxi-Prep:1) Dilute overnight culture 1:10 into LB or 2xYT broth with 100 µg/ml ampicillin.
Grow at 37°C until OD600 reaches 0.6-1.0. It takes about 1-3 hours.
2) Add IPTG to final 0.5 mM, continue growth for 2-3 hours.
3) Spin down cells; freeze at -70°C for 5 min.; thaw in 1:100 to 1:50 culture volume of
PBS containing 1 mM PMSF, 10 mM β-mercaptoethanol, 0.2 mg/ml lysozyme. Leaveat 4°C for 20 min..

150

GST fusions for antibody production4) Add Triton X-100 to 1%; sonicate cells for 30-60" (do not over-sonicate; it may cause
denaturation. Multiple bands will be seen on the SDS gel because of the associationof denatured GST fusion protein with other proteins.)5) Spin 10,000 x g for 5 min. at 4°C.
6) Mix supernatant with 1 ml 50% glutathione-agarose beads (For 500 ml culture) ; leaveat room temperature for 10 min., while slowly shaking; collect beads at 500 x g;
wash 3x 50 ml PBS with 1% Triton X-100; elute fusion protein 2x 2 min. with 1 beadvolume with freshly made 5 mM reduced glutathione in 50 mM Tris-Cl pH 8.0 buffer.
•50% Glutathione-agarose beads (Sulphur linkage, Sigma G-4510) is preswollen in PBS +1% Triton X-100, washed twice and kept at 4°C.
•Beads can be reused after washing in 3 M NaCl.
•Smaller fusion proteins (50kD or less) tend to give better yields; it is also best to avoid
regions of th gene that encode hydrophobic peptides.
Reference:
Smith, D. B. and Johnson, K. S. Single-step purification of polypeptides expressed in
Escherichia coli as fusions with glutathione S-transferase. Gene 67:31-40, 1988.

151

Don RioSOLID PHASE RADIOIMMUNE ASSAY FOR IgG
1)Apply antigen (usually 0.1-1 mg) into polyvinyl chloride well of microtiter plate(96-well) in 50 ml final volume in PBS, pH 7.4 (Costar vinyl assay plates, cat. no.
2596).

2)
3)

Allow solution to evaporate (RT overnight, or 37°C incubator for 1-2 h).
Apply 150 ml 5% BSA, PBS, pH 7.4. (PBS = 0.01M NaPO4 pH 7.4, 0.1 M NaCl).

4)

Replace tray lid; place at 37°C for two hours.

5)

Remove BSA-PBS blocking solution.

6)

Apply antibody in 50 µl final volume, diluting in PBS, pH 7.4 if necessary.
Unconcentrated hybridoma supernatants are used without dilution.

7)

Replace tray lid; place at 37 °C for one hour.

8)

Remove antibody solution.

9)

Wash three times with 1% BSA, PBS, pH 7.4; use about 100-150 µl per wash and10)

discard wash solutions between applications.
Apply 105 cpm [125I] - protein A or appropriate [125I] labeled second antibody
in 50 µl final volume, using PBS, pH 7.4 as diluent.

11)
12)

Replace tray lid; place at 37 °C for one hour.
Remove [125I] solution.

13)

Wash three times with 100-150 µl PBS, pH 7.4.

14)

Aspirate any residual liquid in microtiter wells and wipe the top of the plate dry.

15)

Place plate without lid between two glass plates with X-ray film and screens inplace and use binder clips to clamp.

16)

Autoradiograph with XAR-5 or equivalent film being certain the well bottom isagainst the film; use one intensifying screen; place at -70 °C for 16 hrs orovernight.

PBS:

10 mM NaPO4, pH 7.4
0.1 M NaCl

The pH is very important as the reactions are not efficient below pH 7.0 and are lessefficient at pH 7.0. Therefore, adjust the pH of the PBS and the PBS containing BSA. Store the
BSA solutions at -20 °C.
NOTE: It is necessary to adjust pH of BSA/PBS solutions as pH drops when BSA isdissolved in PBS.

152

Don Rio
Revised by Smadar Admon

PROTEIN GELS - SDS POLYACRYLAMIDE
Stock Solutions: (1) 30% Acrylamide/0.8% Bis(2) 1M Tris, pH 6.8(3) 1M Tris, pH8.8(4) 10% SDS(5) 10% APS(6) Laemmli buffer (sample buffer) (prepare fresh):
For 1 ml:
100 µl glycerol
200µl 10% SDS
125µl 1M Tris, pH 6.850µl BME
0.05% BPB
525µl ddH20(7) Running buffer (for 4 liters): 57.6gr Glycine
12.1gr Tris base
4.0gr SDS
Recipes for gels:% running gel
30%/0.8% AA
ddH20

5%
2.66ml
7.1ml

7%
3.7ml
6.0ml

9%
4.8ml
5.0ml

10%
5.33ml4.43mlAdd: 6ml 1M Tris, pH 8.8160µl 10% SDS
80µl 10% APS
8-15µl Temed
Stacking gel (5%, 5ml): 0.625ml 1M Tris, pH 6.8
0.83ml 30%/0.8% AA
3.46ml ddH2O
50µl 10% SDS
25µl 10% APS
6µl Temed
Staining Gels:
Staining Solution: 10% acetic acid50% methanol
0.25% Coomasie brilliant blue R-250
Destaining Solution: 10% acetic acid20% methanol

153

12%
6.4ml
3.36ml14%
7.46ml2.3ml

16%
8.53ml1.23mlSmadar Admon

WESTERN BLOTS
Stock solutions:(1) Transfer solution (4 liters):
57.6gr glycine
9.7gr Tris base
800ml methanol(2) TBS:

25mM Tris-HCl
150mM NaCl
pH 7.4(3) TTBS :TBS + 0.5% Tween 20(4) Blocking solution:

2% Carnation non-fat dry milk in TBS, pH 7.4.

(5) Ponceau's S stain:

0.5gr Ponceau S + 1ml acetic acid + H2O to 100ml.

Transfer of proteins to nitrocellulose:
After running the acrylamide gel, transfer it to nitrocellulose by electrophoresis. The gel
is put in the transfer apparatus with the nitrocellulose on top of it, facing the positiveelectrode. The transfer should be done in the transfer solution, at 4°C, 20 volts,
overnight, or at 50 volts for 4-5 hours.
After the transfer, folow the procedure below:
1) If you want to check the transfer you could stain the filter with Ponceau S stain(reversible):(a) Stain for 5 min.
(b) Destain in water for 2 min.
(c) Wash off the stain in PBS for 10-30 min.
2) Place nitrocellulose in a Seal-a-Meal bag with blocking solution. Incubate and shakefor at least one hour at room temperature (can be left overnight).
3) Rinse briefly in TTBS.
4) Incubate with primary antibody diluted in blocking solution for 1-2 hours at room
temperature (shaking). Proper amount of antibody is determined empirically.
Supernatants from cells, no dilution - 1:5; Ascites or serum - 1:100 - 1:500.
5) Wash twice for 10 minutes in TTBS.
6) Incubate with secondary antibody diluted in blocking solution at room temperaturefor 1 hr. For AP or HRP conjugates, dilute 1:500 - 1:1000. For 125I-labelled protein Aor anti-IgG use 2 x 105cpm/ml.

154

Western blots7) Wash as in step 5.
8) (a) If using 125I, expose nitrocellulose using Kodak XAR5 film and intensifying
screen overnight at -70°C.
(b) If using AP conjugate, rinse once more for 5 minutes with TBS (no Tween).
Develop blot using the BCIP + NBT substrate system (Bio-Rad kit).

155

Joe Heilig

ELECTROELUTION OF PROTEINS USING ISCO APPARATUS
1)

Run preparative Laemmli SDS/PA gel as desired.

2)

Remove gel from plates, do not fix, and stain for approximately 10 min.

3)

Destain gel.

4)

Rehydrate gel in water for 5-10 minutes.

5)

Cut out region of gel containing band of interest and mince gel slice into ≈1mm
cubes. This can be done conveniently in a weigh boat.

6)

Prepare elution cup by securing a single layer of dialysis membrane beneath each of
the wells with the retaining rings. Place a plastic mesh screen over the collection(smaller) well.

7)

Prepare buffer chamber by placing a piece of Whatman 3mm paper in the divider
slots of each electrode chamber and fill all chambers with 1x Laemmli gel runningbuffer, containing SDS, to approximately 1cm below the top of the central
divider/cup holder. Buffer must not rise over this divider when the elution cups
are placed on it. The paper dividers prevent bubbles generated at electrodes during
elution from collecting under the wells and preventing current flow.
If a very small amount of protein is being eluted (in the pg range) such as might berecovered from a biologically significant source, not a fusion protein, it may be useful to add
protease inhibitors to the elution buffer. For example, add PMSF to 1mM and aprotinin to
1/1000.

8)

Pile minced gel pieces into sample (larger) well. Be careful not to trap air in thebottom of the wells or create air pockets in the mass of minced gel.

9)

Place elution cups in the chamber straddling the center divider. Be certain not totrap air bubbles underneath the wells.

10) Elute in cold-room at 150 V, constant V, for ≈12 hrs. Be certain that there is current
flowing through the chamber. The Coomassie Blue dye will collect in the collection
chamber and provide an indicator of completion of elution.
If no current flow is detected the most likely cause is a bubble either in or under one of thewells. It can sometimes be difficult to see these bubbles so inspect closely, removing the gelslices if necessary.
11) When elution is complete, remove buffer from elution cup with pasteur pipette,remove gel slices from collection well side of cup and remove remaining buffer
down to rim of the collection well.
12) Remove screen and recover eluted protein from collection well with Pipetman orequivalent. The volume of the collection well is 200µl. The collection well can be
washed by pipetting solution up and down, gently so as to prevent foaming, orwith fresh buffer.
13) Dialyse the sample as necessary.

156

Electroelution of prteins using ISCO apparatus14) Clean the apparatus thoroughly.

157

Betsy O'Neill

MAKING FOOTPRINTING EXTRACTS FROM DROSOPHILA HEADSThis is a protocol which I adapted from the standard Tjian lab embryo nuclear
prep. I generally make extracts from 200-500g of adult flies at a time. I harvest theminto disposable 250 ml Corning centrifuge tubes, and store them at -70 until I'm ready touse them (up to a few weeks at least). I separate the heads by placing the tubes into
liquid nitrogen for a few minutes and then shaking the tubes vigorously. This causes
the heads, wings and legs to fall off. I then put this material through two sieves-a
number 25, which retains the abdomen and thorax, and then a number 40 which retains
the heads, but allows the legs and wing fragments to go through. This also requires
vigorous shaking to get good separation. I had an apparatus constructed as
diagrammed below which I place into a multivortexer to acheive separation more easily(I had to do this because my arms got tired before I had acheived good separation).

retains:
thorax /abdomen
heads
legs/wings
misc. fragments

plexigass lid

No.25 sieve
Nitex: same mesh sizeas No. 40 sieve
plexigass bottom

I then place the still frozen heads into about 1ml of high sucrose homogenizing
buffer per 3g of adult flies. I put this mixture through a Yamato continuous flow
homogenizer set at 700 rpm. I collect the flow through and pass it through thehomogenizer another 4-5 times. I periodically examine samples of the flow throughunder a dissecting scope to monitor the efficency of homogenization. This makes it easy
to determine when all of the heads have been broken up. Next, I pass the homogenate
through prewetted (ddH20) miracloth (Calbiochem #475855) placed in a large funnel,
which removes large particulate matter, but allows nuclei to pass through. Nuclei arethen isolated by spinning at 24K for 45 minutes at 4 C in an SW28 rotor. Since the SW28
tubes must be filled completely for the spin, I usually bring the volume of the material
that comes through the miracloth up to something divisible by 40ml, since the tubes
hold about 38ml. I experimented initially with spinning through a sucrose cushion on
this step (hence the SW28 rotor etc.), but had difficulty isolating the nuclei--they seemed
to be at the interface rather than in the pellet as described in the protocol I was trying toadapt, so I eventually settled on the 24K spin. There is generally a large layer of lipid at
the top of the tube after this spin, which I usually scoop off before attempting to pour off
the cytoplasmic fraction.
The pellet (nuclei) from this spin is resuspended in 0.1ml of Buffer A+B per gramof flies. The pellet itself is usually composed of a softer upper layer, the nucei, and ahard, dark lower layer. I usually resuspend only the top layer and discard the lower
layer. Transfer this material into Ti60 tubes. The nuclei are lysed (in the Ti60 tubes, sothat there is no need to transfer the viscous solution containing lysed nuclei) by adding
0.1 volume of 4M ammonium sulfate pH8.0. I usually add this a little at a time, and mix
well, but gently, in order to get even lysis. I then place the tubes on a rotating wheel at 4
C for 20 minutes. Pellet the chromatin by spinning at 35K for 30 minutes in a 60Ti rotor
at 4 C. The supernatant from this spin is a >10% AS cut. The crude extract that I

158

Making footprinting extracts from Drosophila heads

generally work with is a 10-60% AS fraction which I prepare by adding 0.3g of finely
ground ammonium sulfate per ml of extract. We have a coffee bean grinder that I use togrind the AS, but I'm not sure that's necessary. I place the extract in a small beaker in an
ice bath on a stir plate in the cold room, add the AS gradually, and stir for 15 minutes. I
then transfer this material to a Sorval tube, and spin at 15K for 30 minutes. I resuspend
the pellet in 2ml of 0.1M HEMG per 100g flies, and dialyze against 0.1M HEMG until the
conductivity is equal to that of 0.1M HEMG (takes 4-6 hours). This is my final extract
which I aliqout and freeze in liquid nitrogen, and then store at -70 C. The protein
concentration of the extract is usually around 15 mg/ml, using BSA as a standard, orabout 30mg protein per 100g flies. I have experienced quite a bit of variability in yield,
which I think is due mainly to variation in the efficiency of the early steps (i.e.,
harvesting heads). For example, if you don't shake the frozen flies long enough to get all
the heads off, the yield will be lower.
I see good footprints on the rhodopsin promoters using in the range of 10-100 ug
extract in a standard footprinting reaction.
Protocol Summary:1) Harvest heads from adult flies by using appropriate sieves.
2) Place heads into a beaker containing 1 ml homogenizing buffer per 3g adult flies.
3) Homogenize in continuous flow homogenizer at 700 rpm. Collect flow through andrehomogenize until all heads are broken open.
4) Pass homogenate through prewetted miracloth to remove particulate matter.
5) Bring the total volume of this filtrate up to an amount divisible by 40 withhomogenizing buffer, mix well, and transfer to Ultra Clear SW28 tubes (about 38 ml
per tube). Spin at 24K/4 C/45 minutes.
6) Remove upper layer of lipid and pour off supernatant (cyto). Resuspend pellet(nuclei) in 0.1ml Buffer A+B per gram of adult flies.
7) Transfer this material (noting volume) to 60Ti tubes. Lyse nuclei by adding 1/10
volume 4 M ammonium sulfate pH 8.0. Add the AS a little at a time, and mix gently
by capping the tube and inverting it several times. This should cause the mixture to
become quite viscous.
8) Place the tubes on a rotating wheel at 4 C for 20 minutes.
9) Pellet the chromatin by spinning at 35K/4 C/30 minutes.
10) Place the supernatant from this spin (note the volume) into a small beakercontaining a stir bar, and place the beaker in an ice bath on a stir plate in the cold
room. Gradually add 0.3g finely ground ammonium sulfate per ml of supernatant tothe stirring extract. Stir for an additional 15 minutes.
11) Transfer this material to a Sorval tube and spin 15K/4 C/30 minutes.
12) Resuspend the pellet in 2ml 0.1M HEMG per 100g adult flies and dialyze against
0.1M HEMG until the conductivity of the extract equals that of 0.1M HEMG.

159

Making footprinting extracts from Drosophila heads

13) Aliquot a portion of the extract into small aliquots, and then freeze in liquid
nitrogen.
SOLUTIONS
additions per 1 liter
15mM HEPES pH 7.6
15 ml of 1M (pH w/ KOH)
100mM KCl
7.46g
5mM MgCl2
5ml of 1M
just before use add:
1mM DTT
1:1000 dilution of 1M
0.5mM sodium metabisulfite
1:1000 dilution of 95mg/ml
0.1mM PMSF
1:200 dilution of 0.2M
(in EtOH store at 4 C)

Buffer A+B

Homogenizing
10mM HEPES pH 7.6
Buffer
25mM KCl
0.15mM spermidine
0.5mM spermine
1mM EDTA pH 7.9
2M Sucrose
10% glycerol
just before use add:
1mM DTT
0.5mM sodium metabisulfite
0.1mM PMSF
25mM HEPES pH7.6
100mM KCl
12.5mM MgCl2
0.1mM EDTA pH 7.9
10% glycerol
just before use add: (to 2 liters)
1mM DTT
0.5mM sodium metabisulfite
0.1mM PMSF

0.1M HEMG

160

additions per 1 liter
10 ml of 1M
1.87g
174 mg
38 mg
2 ml of 0.5M
685g
100ml
1:1000 dilution of 1M
1:1000 dilution of 95mg/ml
1:200 dilution of 0.2M
additions per 2 liters
50 ml of 1M
14.93g
25 ml of 1M
0.4 ml of 0.5M
200 ml2 ml of 1M
190 mg
1 ml of 0.2M

Making footprinting extracts from Drosophila heads

4M ammonium sulfate
Add 52.8g ammonium sulfate per 100 ml. Adjust the pH with NaOH. Thissolution is almost saturated at room temperature. Filter any undissolved crystals andstore at room temperature.

161

Zhichun Lai

HIGH EFFICIENCY ELECTRO-TRANSFORMATION OF E.COLI
Preparation of Cells:1) Inoculate 1 liter of LB broth with 1/100 volume of a fresh overnight culture.
2) Grow cells at 37°C with vigrous shaking to an OD600 of 0.5 to 1.0 (the best results are
obtained with cells that grow rapidly; the appropriate cell density, therefore, depends
on the strain and growth condition).
3) To harvest, chill the flask on ice for 15 to 30 minutes, and spin in a cold rotor at 4000x
g max for 5 minutes.
4) Remove as much of the supernatant as possible. Resuspend pellets in a total of 1 literof cold water. Spin as in step 3.
5) Resuspend in 0.5 liter of cold water. Spin as in step 3.
6) Resuspend in about 20 ml cold 10% glycerol. Spin as in step 3.
7) Resuspend in a final volume of 2 to 3 ml in cold 10% glycerol. The cell concentration
should be 1-3x1010 cells/ml.
8) The suspension could be frozened in aliquots on dry ice, and stored at -70°C. The cells
are good for at least 6 months under these conditions.
Electro-transformation
1) Gently thaw the cells at room temperature and place them on ice.
2) In a cold, 1.5 ml polypropylene tube, mix 40 µl of the cell suspension with 1-2 µl of
DNA (in either TE or water). Mix well and let sit on ice for about 1 minute.
3) Set the Gene Pulser apparatus at 25 uF and 2.5 kV. Set the Pulse Controller to 200 om.
4) Transfer the mixture of cells and DNA to a cold, 0.2 cm electroporation cuvette, andshake the suspension to the bottom of the cuvette. Place the cuvette in a chilled safety
chamber slide. Push the slide into the chamber until the cuvette is seated between thecontacts in the base of the chamber.
5) Pulse once at the above settings. This should produce a pulse with a time constant of
4-5 msec. (The field strength will be 12.5 kV/cm.)
6) Remove the cuvette from the chamber and immediately add 1 ml of SOC medium tothe cuvette and quickly resuspend the cells with a pasteur pipette.
7) Transfer the cell suspension to a glass culture tube and incubate at 37°C for 1 hourwith shaking.

162

High efficiency electro-transformation of E. coli

8) Plate on selective medium.
For 100 ml SOC medium:Take 94 ml LB medium which contains:
2% Bacto tryptone
0.5% Bacto yeast extract
10 mM NaCl
Add:

2.5 mM KCl
10 mM MgCl2
10 mM MgSO4
20 mM glucose

1M
1M
1M
10 %

Ref: The Bio-rad Instruction Manaual.

163

0.25 ml
1 ml
1 ml
3.57 mlUlrike Heberlein

BROMODEOXYURIDINE (BrdU or BUdR) INCORPORATION AND
VISUALIZATION
BUdR incorporation is a convenient technique for labelling cells undergoing
DNA synthesis. At low concentrations it will not block cell division and will beincorporated into replicating DNA with little apparent damage.
Preparation of BUdR food and feeding:• Make a BUdR stock solution by dissolving 10 mg of BUdR in 350µl of 50% ethanol(28.6 mg/ml) and store at -20°C (it forms a suspension; be sure to vortex before using
an aliquot). BUdR is quite toxic; handle and dispose of responsibly.
• For in vivo labelling weigh and melt some fly food (regular corn meal/agar food) inthe microwave until just liquified. After letting the food cool a bit add the BUdR
suspension in ethanol to a final concentration of 0.5 mg/ml. For short pulses (2-8 hrs)
the concentration can be increased to 1.0 mg/ml. Return the BUdR-food into the vial
and let solidify. In addition (optional) make up some yeast paste and add BUdR to 0.5
mg/ml final concentration. Put this paste on top of the corn meal/agar food.
• Put larvae of the desired age onto this food. To look at eye discs I found that it isnecessary to pull third instar larvae out of the food in their original vials or bottles. If
larvae are already crawling up the walls they don't feed anymore, and obviously
won't incorporate any BUdR.
BUdR visualization:1) Dissect cephalic complexes in PBS and fix for 30 minutes at room temperature in
Carnoy's fix (60 ml ethanol + 10 ml glacial acetic acid + 30 ml chloroform).
2) Rehydrate by incubating for 3 minutes each in 70%, 50%, 30% ethanol in PBS-TX(1xPBS, 0.3% Triton X-100). Finish by incubation in PBS-TX for 5 minutes on ice.
3) Denature the DNA by incubating for 1 hr in 2N HCl/PBS-TX (1:1 mix of 4N HCl and
PBS-TX). Wash twice, 5 minutes each, in PBS-TX on ice.
4) Incubate for 30 minutes in 10% goat serum in PBS-TX.
5) Incubate overnight at 4°C (I imagine that for eye-discs, in contrast to whole embryos
or larval CNSs, a few hours is sufficient) in the primary antibody (1:200 dilution of
monoclonal anti-BUdR from Becton-Dickinson).
6) Wash in PBS-TX on ice six times for 10 minutes each (again, this might be an
exageration when staining eye discs).
7) Incubate at 4°C for several hours to overnight in a 1:200 dilution of biotinylated goatanti-mouse IgG (Vectastain) in PBS-TX.
8) Wash in PBS-TX on ice six times for 10 minutes each.
9) Incubate for 30 minutes in Vectastain ABC reagent (10µl of reagent A + 10µl ofreagent B + 980µl of PBS).
10) Wash in PBS on ice two times for 10 minutes each,164

Bromodeoxyuridine (BrdU or BUdR) incorporation and visualization

11) React with DAB/H2O2/CoCl2 (100µl 5mg/ml DAB + 900µl PBS + 1µl 3% H2O2 +20µl 1% CoCl2) until stain becomes evident, usually a few minutes.
12) Dehydrate discs by a graded series of 30, 50, 70, 90, 100% ethanol.
13) Mount in DPX.

165

Bromodeoxyuridine (BrdU or BUdR) incorporation and visualization

Melissa Cobb

LAMBDASORB LIQUID PHAGE PREP
1)

Inoculate 200µl cells in a 50 ml sterile tube with either 2 plaques or 2 million pfus.
Incubate for 20 minutes at room temp to allow adsorption. Add 15 mls of L-Brothcontaining 10mM MgSO4 and shake at 37°C until lysis (5-7 hrs). Add 150µl
chloroform and shake an additional 15 min to lyse the remaining cells.

2)

Spin at 3,000 for 5 minutes to pellet the cell debris and transfer the supernatant intoa clean tube. Set aside 100µl as high titer stock. Add 100µl of LambdaSorb placetube on a rotator for 30 minutes at room temp or leave at 4°C overnight and rotate
briefly in the morning.

3)

Spin at 3,000 for 5 minutes. Discard supernatant and drain the tubes well.

4)

Resuspend pellet in 1ml TM10 and transfer to eppendorf tube.

5)

Spin very briefly (30 sec), discard supernatant and resuspend pellet in 500µl TE.

6)

Add 5µl 10%SDS, 10µl 250mM EDTA and incubate at 65-70°C for 10 minutes.

7)

Spin 1 minute to pellet protein debris and transfer the supernatant to a new tube.
Add 50µg/ml of Proteinase K and incubate for 30 minutes at 37°C.

8)

Phenol-Chloroform extract twice. Add 50µl 3M NaOAc and 500µl isopropanol andmix well. A DNA precipitate should form readily but, if not leave at -70°C for 15
min, thaw and centrifuge. If the precipitate is readily visible, centrifuge for only aminute so it won't be so hard to resuspend, otherwise spin for 10 minutes.

9)

Resuspend DNA pellet in ~50µl TE.

NOTE: Some members of the lab find steps 7 and 8 to be unnecessary.

166

Dan Kalderon

SMALL-SCALE PREPARATION OF BACTERIOPHAGE LAMBDA DNA
Growing Up Phage
Plate lysate
Make lambda agarose plates (preferably in NZCYM, although LB+ 10mM
MgSO4 is OK; 1.5% agarose).Add 20µl (gt10) or 200 µl (Charon 4) of single plaqueeluate (from 2ml in TM10+chloroform) to 100µl of indicator bacteria in TM10, mix,
incubate 20' at 37°C and spread on a dry 9cm agarose plate in soft agarose (0.7%
48°C).Incubate for about 12 hr. Optimally the plaques should just touch each otherafter about 8 hr but the yield of phage is reduced less by adding too much than too
little phage. As lysis is normally complete it is best also to have one plate with noadded phage to be sure that the plates could support a good lawn of bacteria. Add 5
ml of TM10 to each plate and elute at room temperature for at least 2 hr (or O/N at4°C) with gentle agitation. Collect eluate, adding a further 2 ml TM10 to maximise
yield. At this point a significant proportion of phage still remains on the plate and canbe eluted with a further, say, 5ml of TM10+gelatin to give high titre phagestocks(stored with a little,0.2%,chloroform at 4°C)
Liquid culture
Plate lysates are preferred to liquid culture because fairly reproducible yields ofphage are usually obtained without accurate determination of phage titre.
Nevertheless , some people find it convenient to grow 50-100ml liquid cultures by
innoculating with 1ml of a C600 O/N per 100ml NZCYM and infecting with 0.5-2 X
107 phage (more with Charon 4 than gt10). Grow for 4-7 hr or until lysis occurs.
Preparation of DNA
1)Spin down bacterial (and other) debris from plate lysates or liquid culture (5000rpm 5' Sorvall, 10 ml polypropylene tubes).

2)

Add RNase A & DNase I each to 1µg/ml to supernatant and incubate at 37°C
for 30'.

3)

Add an equal volume of 20% PEG (polyethylene glycol),2M NaCl in TM10, mix
and stand on ice for at least 1-2 hr.

4)Spin down phage (7000 rpm 20' Sorvall SS34), pour off liquid, spin again for 30
secs and aspirate fluid.

5)

Resuspend pellet thoroughly in 0.5ml TM10 and transfer to a microfuge tube.

6)

Extract with 0.5ml chloroform7)

Add 5µl 10% SDS, 10µl 250mM EDTA pH 8 and incubate at 65-70°C for 10mins.

8)

Add 0.5ml phenol (equilibrated with TE) and shake to mix (but do not vortex
henceforth); centrifuge for 2 min and remove aqueous phase which willprobably drag some interphase along due to high viscosity.

167Small-scale preparation of bacteriophage lambda DNA

9)

Extract with phenol/chloroform/iso-amyl alcohol (25:25:1).

10)

Add 0.5ml isopropanol and mix. A DNA precipitate should form but if not,leave at -70°C for >15 min, thaw and centrifuge. If DNA precipitate is readily
visible spin for only a minute (to help subsequent dissolution), otherwise 10
min. Wash pellet in 70% ethanol, dry and dissolve in 50-200µl TE by gentle
shaking or with the aid of a Gilson. The yield of gt10 recombinants is usually
greater than for Charon 4 but the latter normally gives enough DNA for at least
5 restriction enzyme digests. As a large amount of RNA is normally present in
the DNA preparation it is worth treating with boiled RNase A (at ~100µg/ml)
during or prior to digestion, so that (i) small DNA fragments are not obscuredand (ii) the size of fragments can be more accurately estimated from theirelectrophoretic mobilities. (In the presence of excess RNA, all fragments will
migrate faster than they should).

168

Small-scale preparation of bacteriophage lambda DNA

from Grace Gill via Betsy O'Neill

PREPARATION OF SINGLE STRANDED DNA (for in vitro mutagenesis)
Day 1
1)

Innoculate an ON into LB amp of desired plasmid contained in bacterial strain
CJ236, inserts in LB amp.

Day 2
1)

The next day, transfer 50µl of the overnight into 2 ml of LB amp. Shake at 37°C forabout 3 hours, until OD600 is about 0.7.

2)

Add 25µl of helper phage (109 phage for an moi of about 10). Shake at 37°C for 1
hour. (The helper phage we use is VCS-M13, available from Stratagene.)3)

Dilute 400µl into 10 ml of LB amp kan, and shake at 37°C overnight. (Phage is
kanR)

Day 3
1)

Pellet cells in eppendorfs (1.5ml per tube)/10 min.

2)

Carefully remove only 1 ml of the supernatant to a new set of tubes.

3)

Add 400µl 20% PEG 8000, 2.5M NaCl. Vortex and leave at RT for 15 minutes.

4)

Pellet phage/10 minutes in microfuge. A loose pellet should be visible.

5)

Resuspend each pellet in 100µl TE, and combine into two tubes. Phenol/chloroform
extract and ethanol precipitate ss DNA.

6)

Resuspend each pellet in 100µl TE and run 3µl on a gel.

169

Dan Kalderon

BACTERIOPHAGE LAMBDA LIBRARY SCREENING: SUMMARY
1)

INDICATOR BACTERIA: Grow O/N of C600 or Q358, spin down andresuspend in an equal volume of TM10+gelatin (can be stored for a week at
4°C).

2)

TITRE: Make dilutions of phage library in TM10+ gelatin such that 10-100
plaques will form on at least one plate. Mix 100µl of indicator bacteria with anappropriate volume (10-300µl) of phage dilution, incubate for 20 min at 37°C,
add 3ml top agar at 48°C, mix by gentle vortexing or inversion and pour evenly
on a small (9cm) lambda plate that has been dried well (4-12 hr at 37°C, 2-4 hr at
42°C if fresh). Incubate at 37°C for >8hr and count plaques.

3)

PLATING LIBRARY: To 0.3 ml indicator bacteria, add appropriate amount ofphage library stock or dilution (~5-10,000 per plate for genomic, 20-50,000 forcDNA), mix, incubate 20 min at 37°C, add 8ml top (0.7%) agarose at 48°C, mix
and plate on well dried 15cm plates. Incubate at 37°C for 8-16 hr (cDNAs in
lambda gt10 grow much faster than genomic DNAs in Charon 4).

4)

LIFTS: Cool plates (>1 hr at 4°C), label nitrocellulose circles with plate names,
layer smoothly on agarose surface, stab holes through filter and agar withsyringe needle (narrow guage for accuracy) dipped in water-insoluble ink andpeel filter off after 30s (first lift), 60s (second lift), 3 min (third lift), 5min (fourthlift), any time (last lift). Immerse successively in denaturing solution (5-10 min),neutralising solution (10 min) and 2XSSC (to rinse) with occasional agitation.
Store plates in sealed packages at 4°C. Lay filters on 3mm paper to air dry for atleast 30 min before baking 2 hr at 80°C. Remove moisture from vacuum oven as
required.

5)

HYBRIDISATION: Pre-hybridise for 2-24 hr in aqueous buffer (5 or 2XSSCP,
10X Denhardt's, 250µg/ml salmon sperm DNA, (0.1-0.5% SDS)) at 65°C or insame buffer with 15-50% de-ionised formamide at 42°C (no SDS when using
formamide) in bags with ~6 filters each. Add denatured probe (nick-translated,single-stranded, riboprobe, oligonucleotide) to fresh buffer (<106cpm/ml) andreplace pre-hybridisation fluid with ~25ml of this per bag. Incubate at 42°C or
65°C (with shaking if possible) for 8 (nick-translated probe) to 36 hr (singlestranded probes). Save probe (store at -20°C) for re-screen. Wash filters in 0.10.5XSSC, 0.1% SDS at 65°C (high stringency) or 0.5-6XSSC, 0.1% SDS, at 42°C
(low stringency) for a total of 3-4 hr with 3-4 changes. Wrap filters in Saran
wrap (facing a uniform direction for convenience), mark phosphorescent spots
and expose for between 2 and 24 hr.

6)

PLAQUE PURIFICATION Align filters with film and mark syringe needle
positions. Align film with plates and take a small plug (diameter ~2-3mm) of
agarose+agar from around each positive and elute phage in 2ml TM10 (+/gelatin)+2 drops chloroform for >2hr at room temp with shaking. Dilute 1µl
into 2ml TM10 and plate 2-10µl (gt10 libraries) or 20-100µl (Charon 4 libraries)
as before in top agarose on dry small or large plates to give 100-1000 plaques.
Make lifts and repeat screening procedure using same probe (boiled for 5-10
min before use) and pick isolated positives, or purify a further round if
necessary.

170

Bacteriophage lambda library screening: summary

7)

RE-USE OF FILTERS: If filters are kept in Saran wrap at 4°C they may be reprobed after pre-hybridising, probably any time within several weeks (butcheck the status of the plates). As the position of positives is already known it
is not really necessary to boil the first probe off.
SCREENING BACTERIOPHAGE LAMBDA LIBRARIES(A) LIBRARIES
There exist a number of bacteriophage lambda libraries carrying genomic DNA
sequences or cDNA copies of poly A+ RNA isolated from various stages ofdevelopment or different body parts of D. melanogaster.
Considerations in Choice of Library:(1) REPRESENTATIONThe Maniatis (Maniatis et al., 1978) and EMBL (for vector see Frischauf et al., 1983;
Oregon R library, Pirrotta et al., 1983; Canlon S library, V. Pirrotta, unpublished)libraries were made by sonication and partial Sau3A digestion, respectively, of highmolecular weight DNA. The DNA content of each library (6x105 x16 kb ; Maniatis) far
exceeds (1.65x105 kb) that of the Drosophila genome. Theoretically, therefore, thegenome should be fully represented by each of these libraries, with the possible
exception of DNA that is not readily extracted, DNA that has an unusual structure(e.g. heterochromatic) and DNA that contains sequences which cannot be propagatedin rec A+ hosts due to recombination or "poisoning". Practical experience has
confirmed these expectations. It has been possible not only to clone a large number of
specific genes but also to establish an extensive series of overlapping phage inchromosomal "walks" in either of these two phage libraries. Difficulties in cloning
have been encountered for sequences that include a long inverted repeat (head to head
defective P elements at the sn locus). Less severe differences in the ability ofrecombinant phages to grow have been inferred from under- and overrepresentationof particular characterised phage. This effect is rarely important but may beminimised by the use of unamplified libraries.
The lambda EMBL vectors tolerate a large insert (23 kb) and recombinants
apparently grow slightly more vigorously than Charon 4 recombinants. These benefits
are to be weighed against the greater body of gene cloning history with the Maniatis
library and the more random nature of the latter library (from sheared DNA as
opposed to partial Mbo I digestion).
The representation of particular sequences in cDNA libraries is subject to many
more variables than pertain to genomic libraries. Amongst these are the number ofphage with inserts of reasonable size, the abundance of the specific RNA in thesource material for the library, the possible absence of a polyA tract, the possible
presence of strong stop sequences for reverse transcriptase and the underrepresentation of sequences derived from the 5' end of the RNA. It is therefore
strongly recommended that the initial isolation of any particular gene should beattempted using genomic rather than cDNA libraries even if the proportional
representation of the sequence is expected to be higher in the latter library. (Of course

171Bacteriophage lambda library screening: summary

this does not apply when the use of specific cDNA libraries is a crucial step in the
protocol used to identify a particular gene).
The expected frequency of occurrence of a specific cDNA in a library can beroughly predicted from the abundance of the corresponding RNA in the source forthe library. As most cDNA libraries include at least 105-106 and up to 5x107
independent recombinants it should be possible to isolate cDNAs for RNAs ofabundance down to about 0.0001% (assuming that the majority of packaged phage
include bona fide inserts of reasonable length).
(2) LENGTH OF INSERTS
For genomic DNA libraries the constraints of packaging limit the size of inserts to
between 15 and 23 kb.
For cDNA libraries the size of inserts depends upon a large number ofexperimental variables and is thus best judged a posteriori. However, it should be
noted that the method of construction can influence the quality of inserts in a
predictable fashion. (i) The use of hairpin formation to prime the second cDNA strand
rather than G-C tailing makes the presence of full-length or nearly intact 5' ends very
unlikely (G-C tailing is now generally used). (ii) Most cDNA libraries include EcoRI
linkers at each end. Thus, methylation of internal EcoRI sites is required before
addition of linkers to prevent internal cleavage. Although treated with EcoRI
methylase during construction, most libraries nevertheless contain some (~20%)
recombinants that have been cleaved at internal EcoRI sites. A large proportion of
cDNAs that have been examined have additional or 3' derived sequences of 20-30 bp
at their 5' ends. It is not clear if this is a general problem or whether it is specific tothose libraries that have been examined.
PLATING OUT BACTERIOPHAGE LAMBDA LIBRARIES
1. Number of plaques to screena. Genomic library
The number of recombinants of average insert length 16.5 kb that must be
screened to detect a particular sequence in a Drosophila genomic library is given in thefollowing Table:
Number ofrecombinants screened

Probability of finding atleast one phaage

5,000
10,00020,00040,00080,000120,000

39%
63%
86%
98%
99%
100 - 6 x 10-4%

Probable number ofphage containing target
sequence
0.5
1
2
4
8
12

Although the main priority is to isolate at least one recombinant containing the"required sequence" the isolation of several in the first screen is helpful in that (i) it
aids restriction mapping of the region (ii) it will often delimit the location of the172Bacteriophage lambda library screening: summary

desired sequence as a relatively small region of overlap common to the different
lambda phage and (iii) sufficient DNA may be cloned immediately to include all
relevant sequences both 5' and 3' of the gene.
Thus, a genomic library is generally screened by plating out 6-12 15 cm plates,
each with 5-10,000 phage.
b. cDNA library
As the number of cDNAs in a library for a given mRNA species cannot be
accurately predicted, it is best to screen as large a number of recombinants asreasonable. The maximum number of phage that can be plated on one 15 cm plate andstill give a reasonable hybridisation signal is in the order of 20-50,000. Theoretically,
any number of plates can be screened simultaneously. However, for an amplified
library, the number of new (as opposed to clonally related) recombinants being
screened is greatly diminished as the number of plaques being screened approaches
the complexity of the library. Hence it is usually better to screen several libraries up toa fraction (1/3 or 1/2) of their complexity than to screen the equivalent number ofplaques from just one library.
1)

CELLS : Make a saturated culture of the appropriate bacterial strain ( C600 or
Q358 ) in O/N broth + 0.2% maltose. The culture may be grown from single
colonies but can conveniently be grown in only a few hours from (0.1-1 ml)
aliquots of frozen cells. The saturated culture can be stored at 4°C for at least a
week but is best stored after pelleting the cells and resuspending in an
approximately equal volume of TM 10 + gelatin. This avoids any potential
problem of phage destabilisation during absorption due to absence of Mg2+ (O/N broth contains no Mg2+ )2)

PHAGE : Dilute the bacteriophage lambda stock in TM10 (+/- gelatin) so thatthe required number of phage per plate (5-50,000) can be measured accurately(10 µl- 0.5 ml). The titre of the phage should first be accurately measured bymaking dilutions in TM10+gelatin and plating as below. Titres are generally
stable for phage stored at 4°C inTM10 and chloroform.

3)

Mix 2-500 µl of the indicator bacteria in TM10 (~4-20x108 cells; 1 OD600 = 8 X
108 cells/ml) with the appropriate volume of phage dilution. When plating
phage at very high density it is best to use slightly more cells (500 µl of 3xconcentrated cells) so that small plaques form, allowing a more evenrepresentation of phage.

4)

Incubate for 15-20 min at 37°C

5)

Add 8 ml top agarose at 48°C to the mixture of cells and phage. Mix byinversion or gentle vortexing and pour evenly on a 15 cm lambda agarose plate.
Even spreading of top agarose is facilitated if plates are warm (37°C, 42°C).
Allow ~10 mins. to set.

6)

Incubate inverted at 37°C for 8-16 hrs.

IMPORTANT POINTS

173

Bacteriophage lambda library screening: summary(i)

Check for contamination of solutions or bacteria with phage by plating one plate
as for the others but with no added phage.

(ii)

All solutions and media in which phage are stored, mixed or incubated must
contain Mg2+ (~10 mM) otherwise the bacteriophage particles fall apart.This
may be seen as a reduction in titre (possibly down to zero) and can give rise toplaques of variable size.

(iii)

Top agarose (at the right concentration) must be used for plating (as opposed toagar) to minimise the chance of peeling this layer off when taking lifts.

(iv)

Top agarose should not be too hot (>50°C) otherwise small plaques in reduced
titre result.

(v)

Plates must be very dry (no gloss and preferably with ridges) to prevent
problems when taking lifts.

MAKING LIFTS FROM PLATED-OUT LIBRARIES
Multiple lifts from one plate :By leaving the nitrocellulose filters on the plate for successively increasing periodsof time it is possible to make several lifts from one plate that contain reasonably
similar amounts of phage. Duplicates are to be recommended whenever the numberof filters to be screened is not already excessive and particularly in screens at low
stringency where spurious signals (not due to DNA-DNA hybridisation) may appear.
For the times listed below for making four lifts, the first two lifts usually remove
considerably more phage than the third and fourth. It is not possible to reduce thecontact time for the first lift if the plates were originally very dry as the filters require
some time to become evenly wetted.
1)

Cool the plates down to 4°C (>1 hr, exposed in the cold room) so that top
agarose surface is as hard as possible.

2)

Label nitrocellulose circles according to plate designations in non water-solubleink (most ball-points are OK).

3)

Layer individual filters smoothly on the surface of the plates making contactfrom the centre outwards. If the plates were very dry, a gloved hand will be
required to ensure that the entire filter wets in a reasonable time.

4)

With a syringe needle dipped in water-insoluble ink mark at least three (five isbetter) asymmmetrically disposed spots on the filter by stabbing verticallythrough to the agar.The spots should be easily visible but fine enough to allow
accurate alignment of filter and plate (25G5/8 needle is good). Peel the filter off
from one edge and immerse, plaque side up in denaturing solution. Whenmaking multiple lifts peel the filter off after fixed periods of time (after makingalignment spots):
Lift 1 - 30 sec
Lift 2 1 min
Lift 3 3 min
Lift 4 5 min

174

Bacteriophage lambda library screening: summary

To maximise signal strength, leave the filter on the top agarose for as long as
possible for the last lift (some phage will always remain).
5)

Agitate denaturing solution occasionally to ensure filters are being wetted.

6)Transfer filters in same orientation after 5-10 mins. (excess is better than too
little time) to neutralising solution. Leave for 10 mins. with some agitation toallow mixing of solutions at the surface of the filters.

7)

Transfer to 2xSSC to rinse off high salt solutions (2-3 min with agitation or
longer if convenient).

8)

Lay filters, DNA side up on 3 mm paper to air dry for at least 30 min.

9)

Sandwich the filters between 3mm or other (Blot block) paper and bake in thevacuum oven for 2 hr at 80°C. No matter how long the filters were air dried they
will retain a considerable amount of moisture. For the sake of the vacuum
pump, seal off the evacuated chamber and wipe off condensation on the door
intermittently.

10)

The filters can now be stored for several days or weeks or used immediately.
The plates from which the lifts were taken should be stored at 4°C, sealed in aplastic bag and labelled. When using nitrocellulose filters straight from the pack(not sterile) it is common to see some contaminant bacteria on the plates after a
few days. However, such contaminants are generally lost rather than amplified
on subsequent re-screening and amplification of phage.

IMPORTANT POINTS
1)

Filters should peel off easily from the top agarose without removing any
agarose. If some very small pieces of agarose stick they should be removed
while in the denaturing solution. If a lot of the top agarose peels off, stop. Even
after extensive cleaning it is likely that there will be a high hybridisation
background and no guarantee of an even signal. Repeat plating out, checking
that plates are very dry, top agarOSE is of correct composition (should set at~45°C), plates are chilled before lifts are made (and that nitrocellulose wets
easily).

2)

If filters are not neutralised for long enough they will turn yellow and become
brittle upon baking. Whether this affects anything other than the ease ofhandling the filters is debatable.

HYBRIDISATION:
PROBES
1. Nick-translated or random primed plasmid DNA or fragments
2. Single-stranded probes from ss phage or cDNAs
3. RNA probes
4. Oligonucleotide probes

175Bacteriophage lambda library screening: summary

The choice of probe is sometimes dictated by the nature of the experiment, e.g.
screening for genes that are differentially expressed (cDNA) or finding genes fromamino acid sequences (oligonucleotides). However, when there is a choice thefollowing advantages and disadvantages of each type of probe should be considered.
1a)Nick-translated or random primed plasmid DNA: Easiest probe to makereproducibly sufficiently hot for library screening. Do not use if libraries arecontaminated with plasmid sequences.

1b)

Nick-translated or random primed fragments: Although the Drosophila
genomic libraries do not contain sequences highly related to commonly used
plasmids there is some benefit to using fragments as opposed to plasmids toscreen at low stringency. Background hybridisation is generally lower (perhaps
due to impurities in plasmid DNA preps.) and the possibility that a strong
signal is due to plasmid cross-homology is immediately eliminated. Perhaps
surprisingly, the use of fragments purified only once (twice is OK) on an
agarose gel does not eliminate strong hybridisation to plasmid-containing phagein contaminated libraries.

2)

Single-stranded probes: Presumably because all radioactive material added tothe hybridisation is as a defined DNA molecule, the use of M13 derived singlestranded probes gives the cleanest signals. Such probes can also be made ofhigher specific activity than nick-translated probes. However, it is necessary tostart off with a lot of radioactive dNTPs to incorporate the same amount ofradioactivity into single-stranded as nick-translated probes (because in the
nucleotide excess that is used to generate probes of maximal specific activity, a
lot of hot dNTPs are incorporated into vector rather than probe DNA). Hence,starting with the same amount of radioactive dNTPs the final signal achieved(under equivalent conditions) is generally lower (up to 5-fold) using singlestranded probes but much cleaner. The use of single-stranded probes istherefore highly recommended for low stringency hybridisations (sensitivity canbe increased by scaling up from the standard protocol and by hybridising for
longer than for nick-translated probes).
M13-derived "double-stranded" probes can be made as for single-stranded
probes but omitting the fragment purification step and can be used as a
substitute for nick-translated probes. However, (i) if anything, the cold strand
will be in excess over the hot strand; (ii) the probe will be long (which in somepeople's view is not desirable); and (iii) the probe cannot be used at low
stringency (certainly for the Maniatis library) as ~100 signals per 5000 plaques
are seen, presumably as a result of sequences cross-homologous to M13.

3)

RNA probes: RNA probes have not been widely used for screening libraries. It
is straightforward to make very hot probes and backgrounds appear to belower than for nick-translated probes. However, the increased stability of
RNA/RNA and RNA/DNA hybrids relative to DNA/DNA hybrids can lead totwo problems. First, some regions (or all) of the probe may be unavailable forhybridisation, particularly at low stringency, and second, capricious crosshybridisation may (and has) been seen, requiring hybridisation at several
stringencies to assess the significance of such signals.

4)

The highest theoretical specific activity of a terminally labelled oligonucleotide(~104 Ci/mmol) is only about 3-fold lower than for an average nick-translated

176

Bacteriophage lambda library screening: summary

probe of average length 400 bp (assuming 50% of probe is newly synthesised
from a mixture with 2 hot nucleotides at 300 Ci/mmol).
Thus oligonucleotide hybridisation is potentially of similar sensitivity to standard
screens with nick-translated probes. This should allow for the use of degenerateoligonucleotides.
The appropriate conditions to use for the hybridisation of oligonucleotides can beworked out from the following formula (quoted in Meinkoth & Wahl 1984)
Td = 4(G+C) + 2(A+T)
where Td is the temperature at which 50% of duplexes dissociate and A,G,C,T are thenumbers of the corresponding bases in the oligonucleotide. Any mis-match leads to a
reduction of about 5°C in Td. Hybridisation is normally conducted at 5°C below Td.
In practice oligonucleotides of 30 bases with 2-4 mis-matches have successfully
been used by hybridising in 7XSSCP at 37°C and washing in 7XSSCP at 42°C.
PRE-HYBRIDISATION
Filters should be pre-hybridised for as long as possible but 2-4 hrs. is usually
sufficient. As the conditions for low stringency hybridisation allow more non-specificattachment of probe to the filters, 12-24 hr pre-hybridisation is suggested in thisinstance. It is possible to pre-hybridise in one solution and hybridise in another but
logic would suggest that it is best to use the same composition of solution in the twocases. Thus, filters should be pre-hybridised in aqueous solution (see below) at 65°C
or, for low stringency in formamide solutions at 42°C in the same bags to be used forhybridisation. Six filters per bag is certainly not excessive. Large volumes (~50 ml per
bag with 6 filters) should be used for pre-hybridisation.
HYBRIDISATION
a. High stringency
Hybridisation will only take place with perfect or near perfect homologues at 65°C
in the following solution
2X or 5X SSCP
10X Denhardt's
250 µg/ml herring/salmon sperm DNA (should be boiled immediately prior to
use to denature)(0.1-0.5% SDS) - optional
Using 250 ng nick-translated probe (~10-25 X 106 cpm) in 25 ml in each bag with 6
filters, 8hr is probably sufficient (12-16 hr is generally used) for hybridisation toapproach completion. (Theoretically, a probe of 2 kb should be 50% renatured by 8hr). Under these conditions a signal should be observable in 4-12 hr (at room temp.Kodak XAR-5). When using single-stranded probes (generallly at lower concentration

177

Bacteriophage lambda library screening: summary

than nick-translated probes) longer hybridisation times (24-36 hr) should improve thesignal but are not necessary.
b. Low stringency
Although it is possible to screen libraries simultaneously at several stringencies(but take care that all lifts have roughly equal amounts of DNA), it is best either toascertain the optimal stringency for screening by performing genomic Southerns at avariety of stringencies or to use the lowest possible stringency under which signals
stand out relative to background. Another strategy, that of hybridising at very low
stringency and washing at successively increasing stringencies is not recommended as(i) it takes a long time and uses a lot of film; (ii) sometimes probe becomes irreversibly
stuck to filters even though they are being kept ostensibly wet throughout; and (iii)
this procedure does not allow for optimal conditions of hybridisation.
In 5 X SSCP
10 X Denhardt's
250 µg/ml salmon sperm DNA
25% formamide at 42°C, background can be consistently as low as for highstringency hybridisation; 20% formamide is generally also clean but in 15% formamide
at 42°C (and appropriate washing conditions) it is very hard to get clean filters. Under
the latter conditions, probes that recognise essentially unique sequences in the genome
will illuminate a smear of DNA on a genomic Southern that in sum exceeds specifichybridisation.
Each 1% formamide lowers the Tm of a hetero- (or homo-) duplex by between 0.6
and 0.7°C (McConaughy et al., 1969; Meinboth and Wahl, 1983). Thus a range of 1550% formamide at 42°C covers a Tm range of at least 21°C, the upper limit
representing standard conditions for high stringency hybridisation. The Tm ofimperfectly matched duplexes is apparently decreased by about 1°C per 1% in thenumber of mis-matched base pairs (Bonner et al., 1973).
The plaques containing DNA with the best homology to the probe will always
give the strongest signals. However, this signal will decline (and background will
increase) if the stringency becomes too low. Thus, it is best to screen at the highest
stringency at which a signal can be clearly seen on a genomic Southern. The best
general conditions for low stringency hybridisation are to use a single-stranded probe
and to hybridise for 20-36 hr. Although on theoretical grounds it may seem best to
substitute salmon sperm DNA with e.g. E.coli DNA when using probes derived from(for example) a mammalian source at low stringency, this has not been necessary inpractice.
WASHING
After hybridisation, cut a corner off the bag, pour off the fluid into a 15/50 ml
plastic tube (to be used for the re-screen) and immediately cut the bag open andimmerse filters in wash solution.The conditions of washing have to be adjusted to thehybridisation conditions, taking into account that hybridisation is optimal at 25°C
below Tm whereas hybrids, once formed, are stable under conditions muchcloser to the Tm. Thus the following set of conditions may be used :178

Bacteriophage lambda library screening: summary

Hybridization%

Washing

XXSCP

FORM

Th

Tm

Tm-Th

∂Tm=

/°C

/°C

-25/°C

%mis

XSSC

Tw

Tm

Tm-Tw

/°C

/°C

-10°C

0

2

65

91.3

1.3

2.6

0.2

65

72.8

-2.2

0

2

65

91.3

1.3

2.6

0.5

65

80.1

5.1

0

5

65

98.6

8.6

17.2

2

65

91.3

16.3

40

5

42

69.8

2.8

5.6

1

65

85.7

10.7

30

5

42

77.0

10.0

20.0

0.2

42

72.8

20.8

25

5

42

80.6

13.6

27.2

0.5

42

80.1

28.1

20

5

42

84.2

17.2

34.4

2

42

91.3

39.3

15

5

42

87.8

20.8

41.6

5

42

98.6

46.6

0

5

42

98.6

31.6

63.2

5

20

98.6

68.6

NOTE
The above table is based on the following equations and assumptions.
(i)

The Tm of a perfect homoduplex in 1XSSC is given by
Tm = 69.3 + 0.41(%G+C)

Marmur & Doty (1962)

For most vertebrates and Drosophila the G-C content of the genome is about
40%, so that the average Tm at 1XSSC is 85.7°C.
(ii)

The dependence of Tm on ionic strength, µ is given by(Tm)µ2 - (Tm)µ1 = 18.5 log10 (µ2/µ1)(iii)

The Tm of duplexes is lowered by 0.72°C for every 1% of formamide in solution(Bonner et al. 1973).

(iv)

The optimal temperature for hybridisation is 25°C below Tm. Hence the fifth
column should be close to zero for perfect duplexes. It has been found (Bonneret al. 1973) that optimal rates of reassociation for two imperfectly matchedstrands occurs at a temperature that is lower than for a perfect duplex byapproximately half the difference between the two Tms. Thus conditions thatgive a (Tm-Th-25)=n are ideal for a heteroduplex in which the Tm is 2n lower
than for a homoduplex (∂Tm= 2 X (Tm-Th-25)).

(v)

The Tm of a heteroduplex that contains n% mis-matches is lowered roughly n°C
relative to a homoduplex (Bonner et al. 1973).

179

Bacteriophage lambda library screening: summary(vi)

The washing conditions given are such that the Tm of a perfect duplex isroughly 10°C higher than the washing temperature (Tw) and correspondingly
higher for imperfectly matched heteroduplexes (columns 6 and 10 should beroughly the same)(vii) All figures given are derived from solution hybridisation data. These conditions
have been succesfully used in practice (actually, with wash conditions shifted
up by one position on the table).
All washing solutions normally include 0.1-0.5% SDS
After the last wash, replace with a little fresh washing solution; take filters fromthis clean solution and wrap between sheets of Saran wrap while still moist.
This allows subsequent washing at higher stringency or removal of probe prior
to re-use of filters. Put phosphorescent markers on Saran wrap for alignment
and expose.
Important Points:
1)

Formamide used in hybridisations should be of good quality: freshly de-ionised
or stored frozen.

2)

As it can be a relatively long time before the reason for a hybridisation signal iselucidated, it is worthwhile clarifying some of the possibilities in the initial
screen by(a) using a probe that does not include any vector sequence.
(b) including a genomic Southern in the hybridisation (particularly at low
stringency, so that the number of different DNAs being illuminated can be
judged).
(c) using two or more different probes either to circumvent possible vector
contamination or to define more precisely the nature of the hybridising DNA.

3)

Background
The inclusion of a genomic Southern should help to distinguish the source of the
problem.
(a) Filters: Any agarose sticking to the filters while making lifts will give rise tobackground. Batches of filters can vary but this has not often proved to be a
problem. It is also not clear that yellow filters (due to incomplete
neutralisation) lead to high background; there is a consensus that rinsing thefilters before baking is important.
(b) Probe: Nick-translated probes used at high radioactive concentration(106cpm/ml-do not exceed this) will always give some (uniform)background, but signals are also strong and very easily visible. The
parameter most strongly favoured for adjustment to optimise signal/noise is
the DNase I concentration used in nick-translation. Single-stranded probes ingeneral do not give any significant background (when made under standard

180

Bacteriophage lambda library screening: summary

conditions); rather the only problem here is the strength of the signal. If thesignal is very low (or zero) make sure the probe really was single-stranded.
(c) Hybridisation conditions: As the kinetics of the hybridisation are generally
not a problem it is best to use large hybridisation volumes to allow easy flow
over the (multiple) filters and to keep the concentration of counts below
106/ml. Also, use plenty of pre-hybridisation fluid as there is a large surface
area to coat and pre-hybridise for as long as possible, particularly at low
stringency.
PICKING POSITIVES & PLAQUE PURIFYING
1)

Align autoradiographs with filters and mark the position of the syringe needle
holes on the film.

2)Mark positives with unique labels, so that after plaque purification it is possible
to return to any plaques that did not check out as expected. To distinguishpositives from background due to cassette contamination, cosmic rays andbubbles on the original plate the following criteria can be used :(i) Shape: the source of radioactivity should be diffuse (as opposed to a point
source) and should be roughly circular. Sometimes plaques will streak whenlifts are taken giving a comet (a circular spot and a tail, not necessarily
connected); this is a clear indication of a real positive. Duplicates help toidentify the original position of the plaque but, irrespective of relative intensity,
the circular region of a comet signal is the place to pick.
(ii) Intensity: Partial overlap between probe and recombinant phage DNA as
well as variable plaque size means that wide variation in signal intensity ispossible. Nonetheless, intensity is the best available guide to distinguishpositives from background and also for low stringency hybridisation to
distinguish between the quality of homology between the probe and various
phage.
(iii) Clustering of positives is a clear indication of artifactual signals.
(iv) Number: Expected numbers of positives can be worked out for genomic
libraries and, less reliably, for cDNA libraries. If there are an excessive numberof positives, consider the possibilities that (i) the probe contains some repetitive
sequences or (ii) the probe includes some "vector sequences" that are alsopresent in the library (pUC, pBR, polylinker, ß-galactosidase sequences).

3)

Align syringe needle marks on plates with their positions on the films and takea plug of agarose (down to the bottom of the plate) that overlies each positive.
Because of the inaccuracy in lining up the orientation marks and because thefilters generally contract a little during hybridisation and washing (if not preautoclaved) it is best to pick a small area around the positive such that you are
confident that the positive must have been picked even given a maximal error inalignment. This can be done by using the wider end of a Pasteur pipette but isbetter done with multiple stabs of the other end to pick a (roughly) circular areaof about 1/3-1/4 the size of the large end.

181

Bacteriophage lambda library screening: summary

4)

Elute the phage from the agar/agarose plug in 2ml of TM10 (+/-gelatin) with 12 drops of chloroform for at least 2 hr. Elution for less than 2 hr will lead to
uneven titres. The titre of phage in the eluate is somewhat variable, depending
on plating conditions, the number of lifts taken and (reproducibly) the type ofbacteriophage lambda.
For the Maniatis library each plaque should yield a titre of 106-107 pfu. For
cDNA libraries in gt10 the titre is 5-10 fold higher. The method of plaquepicking described above removes 10-20 plaques from plates with 10-20,000
plaques. Thus, the expected number of infectious phage in the eluate will be in
the range 1X107-1X108 (Maniatis) and 1X108-1X109(most cDNA libraries). It
should only be necessary to plate out about 100 phage to find several positives;
however, because of variability in the growth rate of phage and the possibility
that the required plaque was initially comparitively small on a crowded plate it
is often worth plating out a large number (500-1000) to be certain of not losing a
positive. In the latter case a large plate must be used. Thus, the following
dilutions are recommended for plaque purification with an error margin that
may lead to overcrowding rather than a loss of a positive signal. Dilutions are
made in TM10.
Maniatis:[Plug in 2ml] 1µl→[2ml] Plate 50-100µl → ~500 plaques
gt10 libraries:[Plug in 2ml] 1µl→ [2ml] Plate 5-10µl → ~500 plaques
Phage are plated out as for the initial plating but using (1/3) fewer bacteria and
3 ml top agarose if using small rather than large plates. Should the number ofplaques be too great or too small return either to the original eluate or dilution,
both of which should be of stable titre if kept at 4°C (or room temp. for a day).
However, if the titre was initially lower than expected due to incomplete
elution, it will now be higher than "expected" as elution will now be complete.

5)Make lifts, pre-hybridise, hybridise and wash as for the initial screen. The sameprobe can be used for initial and second screen but as most will have renatured
during the first hybridisation it should be boiled for 5-10 min immediately prior
to second use. (Precipitation of, presumably, BSA occurs if boiling is for too
long but this does not affect the results whether additional hybridisation
solution is added or not).

6)

Signals from positives should be more numerous and stronger than in the initial
screen – it is therefore not worthwhile making duplicates for a re-screen if only
one probe is being used. Positive plaques to be amplified should be wellisolated in order to avoid contamination by neighbouring, diffusing phage. The
potential problem of contamination is less serious for cDNAs where, in general,a single EcoRI insert will immediately be sub-cloned than for genomic lambda
DNAs where mapping or use as an in situ hybridisation probe may becomplicated by small amounts of contaminating phage.

7)

Stocks of phage can be kept in TM10+gelatin+0.2% chloroform at 4°C or in 7%
DMSO, TM10+gelatin at -70°C after snap-freezing. Such stocks can be madefrom single plaque eluates (106-107pfu/ml) or, more conveniently if they are to182Bacteriophage lambda library screening: summary

be used as a source for large scale growth of phage, as high titre eluates of platelysates (1010pfu/ml).
ARRAYS
It is sometimes desirable to investigate the nature of a large number of positivelyhybridising phage by further hybridisation before preparing DNA. For this purpose it
is convenient to generate multiple copies of a set of positives. Put 100µl aliquots (induplicate if space permits) of each single plaque eluate in a well of a micro-titre dish.
Pour large plates with indicator bacteria (but no phage) in top agarose. Transfer thephage array to such plates (after the top agarose has set) by dipping the "clonemaster"(an array of metal prongs) in the micro-titre dish and then resting it on the surface ofthe plate. Allow liquid to be absorbed and incubate at 37°C for >8 hr. Make lifts andscreen as described above but expect much stronger signals (and sometimes rings
rather than spots; rings are usually observed using phage in gt10 and can potentiallybe eliminated by dilution of the phage).
RE-USE OF FILTERS
If moist filters (stored between sheets of Saran wrap) and plates are stored at 4°C,
the filters can be re-used several times for reasons of economy and in some cases, suchas chromosome walking, as part of the experimental design.
STRAINS AND MEDIA
C600
Q358

(F-, thi-1, htr-1, LeuB6, lacY1, tonA21, supE44, λ−)(hsdR-R, hsdM+R, supF, ø80r)

Lambda plates

NZCYM or LBM+ 1.5% agar

Top agarose

NZCYM or LBM+ 0.7% agarose

NZCYM per litre

10 g
5g
5g
1g
1.2 g

LBM per litre 10 g

Bacto-tryptone
5g
Bacto yeast extract
10 g
NaCl
1.2 g
MgSO4

NZamineNaClyeast extract
casamino acids
MgSO4

REFERENCES
Bonner, T.I., Brenner, D.J., Neufeld, B.R. and Britten, R.J. (1973) Reduction in the rateof DNA reassociation by sequence divergence
Frischauf, A.M., Lehrach, H., Poustka, A. and Murray, N. (1983) Lambda replacement
vectors carrying polylinker sequences. J. Mol. Biol. 170 827-842

183

Bacteriophage lambda library screening: summary

Maniatis, T., Hardison, R.C., Lacy, E., Lauer, J., O'Connell, C., Quon, D.,Sim, G.K. andEfstradiadis, A. (1978) The isolation of structural genes from libraries of eukaryotic
DNA. Cell 15 687-701
Marmur, J. and Doty, P. (1962) Determination of the base composition of
deoxyribonucleic acid from its thermal denaturation temperature. J.Mol.Biol. 5 109118
Marmur, J. (1961) J.Mol.Biol. 3 208
McConaughy, R.L., Laird, C.D. and McCarthy, B.J.(1969) Nucleic acid reassociation informamide. Biochem. 8 3289-3295
Meinkoth, J. and Wahl, G. (1984) Hybridization of nucleic acids immobilized on solid
supports. Analyt.Bioch. 138 267-284
Pirrotta, V., Hadfield, C. and Pretorius, G.H.J. (1983) Microdissection and cloning of
the white locus and 3B1-3C2 region of the Drosophila X chromosome. EMBO J. 2 927934.

184

Hermann Steller

RAPID SMALL SCALE ISOLATION OF DROSOPHILA DNA AND RNA
DNA:
The following protocol describes the isolation of DNA from adult flies but can be usedequally well to extract DNA from other developmental stages. DNA preparedaccording to that method is readily digested by restriction enzymes and has anaverage size of 40-60 kb. In order to remove RNA contaminating the DNA
preparations DNase free RNase should be included when digesting with restriction
enzymes.
1)

Anesthesize flies with CO2 or ether and put 1-20 flies in an eppendorf tube.
Keep on ice until next step.

2)

Add solution A containing 0.1 M Tris-HCl, pH 9.0; 0.1 M EDTA; 1% SDS and
0.5-1% DEPC (added directly before use) and homogenize gently with a 3 mmdiameter glass or metal rod . Use 100 µl of solution A for extracting DNA from
1-5 flies, 200 µl for 6-10 flies and 500 µl for up to 50 flies. Incubate for 20-30
minutes at 70°C.

3)

Add 14 µl of 8 M potassium acetate for each 100 µl homogenate and leave on ice
for 30 minutes.

4)

Spin in the eppendorf centrifuge at 4°C for 15 minutes. Transfer the supernatant
into a fresh eppendorf tube being careful not to disturb the pellet. If you get
flakes in the supernatant respin.

5)

Precipitate DNA by adding 0.5 volumes of isopropanol at room temperatureand spin for 5 minutes at RT. Wash the pellet carefully with 70% EtOH, respin,
dry and redissolve in 10 (1 fly) to 100 (50 flies) µl TE.

185

Rapid small-scale isolation of Drosophila DNA and RNA

RNA:This is a modification of the method of Chirgwin et al. (Biochemistry 18, 5294-5299,
1979) which can be used for the rapid extraction of total RNA form all developmental
stages of Drosophila.
1)

Homogenize tissue (e.g. 1 fly or 50-100 embryos in 50 µl and up to 200 flies in 1ml) GHCl buffer (7.5 M guanidium hydrochloride; 0.025 M NaOAc, pH 7.0; 5
mM DTT) + 0.5% N-laurylsarcosinate + 0.5% DEPC.

2)Extract once with an equal volume of phenol-chlorororm and separate phasesby centrifugation.

3)

Transfer the aqueous (upper) phase into a new tube and precipitate the RNA byadding 1 µl 1 M acetic acid and 25 µl ethanol for each 50 µl of GHCl-solution.
Leave at -20°C for 3-24 hours. Pellet the RNA by centrifugation for 5 min., 812K.

4)

Remove supernatant as complete as possible and redissolve the pellet in half ofthe original volume (but minimally 50 µl) GHCl buffer.

5)

Reprecipitate the RNA as in step 3. (The reprecipitation serves to remove DNA
which will not precipitate under these conditions).

5a)

optional: reprecipitate the RNA for 1-2 more times by repeating steps no. 3 and4. This is only necessary if large amounts of DNA have to be removed, e.g. whenanalysing the transient expression of DNA injected into embryos.

6)

Wash (and store) the RNA pellet in 100% ethanol, room temperature. For
Northern analysis the RNA is best dissolved directly in Northern sample buffer(e.g. 50% formamide, 2.2 M formaldehyde, 1x MOPS).

186

Mary Mullins

CAT ASSAY: DROSOPHILA TRANSFORMANTS
1.

Freeze flies to be assayed in an Eppendorf tube (1.5 ml) at -80°C for 15 min.

2.

Add l00 µl of 0.25M Tris (pH 7.8) and homogenize.

3.

Freeze the homogenate, thaw, and sonicate (optional) for 1 minute.

4.

Heat at 65°C for 5 minutes. This precipitates most other proteins including
some inhibitor(s) of CAT activity.

5.

Spin in cold (rm. temp O.K.) microfuge for 10 minutes.

6.

Transfer supernatant to new tube. For quantitative assays, do Bradford assays

7.

8.

on all extracts so equal amounts of protein can be used in each assay.
Set up reactions:
Extract: 1-90 µl
0.25M Tris-7.8 + 14C-chloramphenicol(CAM): 30 µl=29 µl Tris + 1 µl
CAM (0.2µCi)
4mM Acetyl CoA (Lithium) : 20µl
0.25M Tris: adjust final volume to 140µl
Incubate at 37°C from l5 minutes to several hours. For good quantitation, a timecourse can be done to determine the linear range of the assay.

9.

Stop the reaction and extract the chloramphenicol by adding 0.5ml ofethyl
acetate. Vortex well and spin 1 minute in microfuge. Remove the top organic
phase and place in new tube.

10.

Dry in Speedvac (about 15 minutes) or overnight in the hood.

11.

Resuspend in about l5 µl ethyl acetate.

12.

Spot onto TLC plates (American Scientific Products, Baker-Flex TLC Silica Gel
1B Plates, 20x20cm, Cat. # 4462).

13.

Run in 95:5 chloroform:methanol for l.5-2.5 hours. in a tightly sealed
chromatography tank.

14.

Dry plate and expose at room temp.

15.

Four spots may be seen. The lowest is the non-acetylated CAM, the next two
spots are two different monoacetylated CAM forms, the top spot ( which may
not be seen depending on the duration of the reaction) is diacetylated CAM. For
quantitative assays, the spots can be cut out and counted in a scintillation

counter.
Materials:•14C-Chloramphenicol(CAM): Amersham, CFA515, 54mCi/mmole

187

Kevin Jones

CHROMOSOME WALKING USING COSMIDS
I will divide this protocol into two sections. The first section will describe thescreening of the cosmid library and isolation of positive colonies. The second section
will describe the endcloning method used to isolate end fragments of the cosmids forchromosome walking.
A. Screening of Cosmid Libraries
1. Preparation of Nitrocellulose Filtersi) Wash the desired number of nitrocellulose filters (we use 82 mm diameter
Schleicher and Schuell) in two changes of distilled water. This removes detergent andother impurities which can retard colony growth and/or reduce the titer.
ii) Stack the filters, separating them with sheets of Whatman 3MM paper. Put two
sheets of blotter paper on each end of the stack and wet the blotter sheets with
distilled water. Package the resulting stack in aluminum foil and autoclave 20
minutes. This helps prevent shrinkage of the filters from occurring during the
hybridization and washing process, which can make aligning signals with colonies
on the master plates difficult.
iii) Number the filters and place face down on 30µg/ml ampicillin plates. Thisequilibrates the filters with the media. After a few minutes, flip the filters.
iv) Plate the titered library at a density of 2-4000 colonies per plate. At 37°C it
typically takes about 10 hr for the colonies to reach the proper size for replica plating.
2. Replica Platingi) Allow colonies to grow to about 0.5 mm in diameter. Larger colonies are more
likely to smear, smaller ones don't always transfer evenly.
ii) Pre-equilibrate filters to be used as replicas as in step 3 above. Up to two
replicas can be reliably made with one master filter. Additional replicas can be madefrom a master after the colonies are re-grown.
iii) Replica plate by setting the pre-equilibrated filters onto a couple sheets of thesterile Whatman 3MM paper (generated in step 2 under "Preparation of
Nitrocellulose Filters"), laying the master filter on top of the replica, and putting yourweight on the filters using a replica plating tool. Use a 25 ga needle to put holes
through the filters while they are stuck together. This will allow alignment of signals
with colonies later. It is a good idea to press on the filters with a pair of forceps nearwhere the holes are being made with the needle to prevent the filters from slipping
against each other.
iv) Grow the colonies back up. The masters usually take 3-5 hr at 37°C, thereplicas anywhere from 4-8 hr. The colonies can then be lysed directly or the cosmid
DNA amplified by placing the filters on 100µg/ml chloramphenicol plates for 16 hr.
Chloramphenicol amplification generally improves signal to noise.

188

Chromosome walking using cosmids

3. Lysis of Colonies, Hybridization and Washingi) Lyse the colonies by laying the filters on blotter saturated with 10% SDS 3
minutes, then 0.5M NaOH/1.5M NaCl 5-10 minutes, followed by 1M Tris/3M NaCl,
pH 7.4 for 5-10 minutes. The SDS step is reputed to sharpen signals.
ii) Lay the filters on top of, and then submerge them in 2xSSCP/0.1% SDS. Wipethe colony debris off the submerged filter gently with a folded Kimwipe. Excessive"polishing" will remove the cosmid DNA, the right amount will reduce background.
Just wipe hard enough to remove the debris.
iii) Bake the filters, separated by sheets of Whatman 3MM, under vacuum at 80
degrees Celsius for 2-4hr. Longer baking times help reduce DNA loss from filters thatare to be repeatedly hybridized, washed, and boiled.
iv) Prehybridize the filters in 3xSSCP/10xDenhardts/250µg/ml carrier
DNA/0.1% SDS. A 1hr prehybridization is usually sufficient, unless an unusuallylarge amount of probe is to be added or the hybridization is to last unusually long.
v) Hybridize the probe to the filters in the same type of buffer as used for the
prehybridization. We usually hybridize nick translated 32P probes at a concentrationof about 2x105 cpm Cherenkov per ml for 12-16 hr.
vi) Wash the filters 5 minutes in 2xSSCP/0.1% SDS at room temperature, then 15
minutes in 2xSSCP/0.1% SDS at 65oC, then in 2xSSCP/0.1% SDS until backgroundcannot be heard on the filters with a monitor. We usually let this last wash go about
2hr. Sandwich the filters between two pieces of Saran wrap and apply to X-ray film.
4. Isolation of positive coloniesi) Align the autoradiogram with the filters and mark the positions of the holes.
ii) Ethanol wipe a piece of cellophane (or something similar). Spot about 0.2 ml of
L-broth onto this in a number of droplets. Lay the master filter (with colonies) onto
the L-broth, set the cellophane on the autoradiogram. Align the holes, sometimes
background on the autoradiogram helps in the alignment. You never thoughtbackground could be useful, right? Get the alignment as precise as possible.
iii) If you're lucky you'll have one good candidate for the signal. If not, pick eachcolony individually with a sterile toothpick and transfer to a plate in a grid pattern. If
there are double colonies, etc. pick these and streak them to get single colonies which
can be put in a grid. Grow up the grid, lift the colonies from the plate using anitrocellulose filter (make holes in the filter with an india ink filled hypodermic
needle before removing the filter to help align it with the plate later) and treat thefilter as above.
iv) Cosmid DNA from the resulting positive colonies can be isolated using a
number of methods. The most rapid I've used is to make a boiling miniprep (seeQuigley and Holmes, Analyt. Biochem. 114, 193 (1981)) of 1.5ml of a saturated culture(I use L-broth with 100µg/ml ampicillin). This generally yields roughly 5-10 µg of
DNA. I've heard that alkaline lysis miniprep methods can give better yields but after
determining that the cosmid is what I want using this miniprep DNA I just go ahead

189

Chromosome walking using cosmids

and grow a 500ml or 1 liter culture and purify the cosmid by alkaline lysis followed
by a CsCl gradient.

B. Isolation of cosmid end fragments by subcloning
The idea here is to isolate cosmid end fragments as rapidly as possible whenchromosome walking. Many cosmid libraries are constructed using vectors whereend fragments from either end of the cosmid can be easily subcloned. For example,
the cosPneo vector has a polylinker containing a BamH1 site flanked on one side by
an Eco R1 site and on the other side by an Xba 1 site. If the cosmid library in thisexample is made by inserting genomic DNA into the Bam H1 site, the two ends of thecosmid can be isolated by intramolecular ligation of complete Xba 1 and Eco R1
digests.
Miniprep DNA is sufficient for this procedure as long as you can digest it tocompletion. Digest 0.5-1.0 µg of cosmid DNA in a 50 µl reaction. Check half of thedigest on a gel to be sure it is complete. Heat kill the enzyme if possible. Otherwise,bring the reaction to 50 µl and add diethylpyrocarbonate (0.5 µl of 10% DEP in
EtOH), heat to 37° for 20 min. Add 2.5 µl 1M Tris (pH 8.0) and heat to 65°C about 10
min. to ensure that the DEP is inactivated. The digest can be run over a P10 spincolumn at this point to ensure high cloning efficiency, but this is usually unnecessary.
In any case, bring the reaction to 100 µl and add ligase buffer and 100 units T4 DNA
ligase. Transform competent cells as usual. The resulting endclones can be hybridized
to cosmids from the previous step in the walk to determine which one extends thewalk. They can also be used for in situ hybridization to polytene chromosomes to getan accurate picture of the progress of the walk.

190

Chromosome walking using cosmids

Tyler Cutforth

DEEP PSEUDOPUPIL IN WHITE- AND RED-EYED FLIESThe deep pseudopupil is an optical phenomenon arising from the regulararrangement of ommatidia in the compound eye and the nature of the rhabdomeres inDiptera, which are open and can transmit light ( N. Franceschini in Information processingin the visual systems of arthropods [ed R. Wehner], pp. 75-82, 1972). By focussing in a plane
below the outer surface of the eye, virtual images of the rhabdomere patterns fromseveral ommatidia are superimposed and the stereotypic trapezoid arrangement ofphotoreceptor cells R1-R7 is apparent. We use the deep pseudopupil to score living flies
for defects in photoreceptor arrangement which may be masked by a normal external
morphology.
Observation of the deep pseudopupil of red-eyed flies requires illumination frombelow (antidromic), using a narrow, bright light source (such as the Magnalite T-150)
with a dissecting microscope and a transparent, elevated stage for the flies. Positioning
the light source is a matter of trial and error, however a general guideline is to place thetips of the light pipes just inside the field of view under low power and angled upwards
to illuminate the centre of the field. The pseudopupil itself is best viewed under highpower (5X), with the anesthetized flies lying on their side, backs toward the observer.
This position permits scoring of the posterodorsal quadrant of the eye, and the lightpipes may need adjustment at this point to maximize the contrast between thepseudopupil and the surrounding eye surface. Other regions of the eye may be scoredby tilting the head, but this is more problematic since slight motions of the hand blur theimage. The deep pseudopupil of white-eyed flies is viewed with illumination fromabove (orthodromic), but in all other respects is similar to that of red-eyed flies.

191

Gerry Rubin

MAKING SEMINAR SLIDES
1. Color Slides
These are quick to make and give accurate color reproduction; good for half-tones
such as autoradiograms. Black and white diagrams have a tendency to come out darkgray on pale yellow, but are certainly usable.
Film:

Kodak Ektrachrome 50(Tungsten)

Shoot:

Using light meter: no filter on camera. Exposure time isaccurate for half tones. For originals having a highpercentage of white area, the light meter will not give thecorrect exposure and you must correct by using a longer
exposure, for example, two stops longer for a typewritten
table (e.g. 1/15 sec. vs. 1/60 sec.). Always shoot a range - onestop on each side of what you judge to be the correct
exposure.

Development:

E6 procedure. Ziba does two runs a day, Monday – Friday:
9:30 - 1 pm and 12 noon - 4 pm.
Presto Prints (on Telegraph) does processing with a two-hourturnaround and is open on weekends. Latest time for having
film developed that day is 3:30p.m.

2. White on blue slides
Good for black and white diagrams. Can also get some additional colors:
Original
Slide
black
white
white
dark blue
dark red
light blue
dark greenpinklight blue
light purple
Film:

Kodak SO-279 veracolor slide film.

Shoot:

A range from f8 - f11 with a 1 sec exposure with our camera
setup. Use HOYA 85B and Nikon Y52 filters.

Development:

C-41 processing to negatives takes about 1 hour at Ag Photo(Shattuck Center across from Andronico's) ~$3/36 exp roll.

3. Computer slides on slide maker
Using a computer program that alows you to save in PICT format (Dreams,
Canvas, MacDraw, but not Word), make your diagram and save a copy as a PICT file.
Follow instructions in Imager manual to print slides on Slidemaker.

192

APPENDIX I: REAGENTS
Acrylamide stock (38%/2%) (Hazardous to your health. Wear gloves. Work in hood.)
76 g acrylamide
4 g methylene bisacrylamide
Make up to 200 ml with DDH2O. Store in dark at 4°. (Wrap bottle with tin foil.)
Amp Plates
5 g NaCl
10 g Bactotryptone
5 g yeast extract
15 g agar
DDH2O to 1 liter
Autoclave
Add 0.3 ml amp stock just before pouring.
Ampicillin
Stock = 250 mg/ml
1 g ampicillin sodium (pharmaceutical grade) in 4 ml sterile DDH2O
Store in aliquots at -20°
10X BU Buffer
70 g Na2HPO4 or 132 g Na2HPO4 • 7 H2O
30 g KH2PO4
DDH2O to 1 liter
Autoclave
Benton and Davis Solutions
Denat.
0.1 N NaOH
1.5 M NaCl
Neut.

0.2 M Tris pH 7.5
pH to 7.5 with HCl

4 g/l or 10 ml of 10N
87.5 g/l
24.2 g/l= ~25 ml conc HCl per 2 l

Optical Grade CsCl Step Grade Sol'n
1.3 g /ml
1.4 g/ml
1.5 g/ml
1.6 g/ml

CsCl
3.1 g
3.9 g
4.5g
5.1 g

TM10
6.9 ml
6.1 ml
5.5 ml
4.9 mlVortex. Should go into sol'n easily.
50 mM CaCl2

193

Appendix 1: Reagents

3.7 g CaCl2 • 2 H2ODissolve in 500 ml DD H2O.
Autoclave.
Casamino Acids for Sinsheimer's
9.9 g casamino acids in 837 ml DDH2O
Autoclave.
(Use 1 liter Bellco bottles.)
Chloramphenicol Stock - 45 mg/ml
9 g chloramphenicol
200 ml EtOH
Store at -20°C. Use 3.5 ml/liter.
20 mM DTT
40 µl 1 M DTT stock sol'n
1960 µl chilled sterile water
Aliquots of 100 µl/Eppendorf tube. Freeze.
Dialysis Bags
sodium carbonate 400g (10% )EDTA 74.4 g (50 mM )H2O to 4 liters
boil 4X
250 mM EDTA pH 8.0
186.1 g Na2 EDTA • 2 H2ODissolve in ~1700 ml DD H2O. Adjust pH with 50% NaOH to pH 8.0 (~28 ml 50% NaOH).
EDTA won't dissolve until the pH is increased. Make up to 2 liters with DD H2O.
Autoclave.
Ficoll/Bromphenol Blue Loading Buffer
6 g Ficoll
43 g H2O
Heat with stirring.
Add pinch of Bromphenol blue
5 ml 250 mM EDTA
Fly Egg Laying Medium (for 18 trays 5" x 5")2225 ml H2O
88 g agar
360 ml molasses

194

Appendix 1: Reagents

Autoclave 30 min.
Add 0.5 ml conc. Tegosept (16 g/150 ml 95% EtOH)
20 ml of ethylacetate, after autoclaving.
Freezing Medium (2X)
3.0 g NaCl
12.6 g K2HPO4 (anhydrous)
0.9 g NaCitrate • 2 H2O
0.18 g MgSO4 •7 H2O or 0.09 g MgSO4
1.8 g (NH4)2SO4
3.6 g KH2PO4 (anhydrous)
88 g (70 ml) glycerol (glycerin)
Dissolve and adjust volume to 1 liter. Autoclave.
Gelatin - 10 mg/ml or 1%
2 g gelatin
200 ml DD H2OHeat and stir to dissolve
Autoclave.
10% Glucose
200 g glucose
Dissolve in 2 liters DD H2O
Autoclave.
Hot Fudge Sauce
1/2 cup butter
3 squares baking chocolate
1-1/2 cups sugar
1/2 cup cocoa

dash of salt
1 cup light cream or evaporated milk
2 teaspoons vanilla

Melt butter and chocolate over low heat. Mix in sugar, cocoa and salt until smooth. Add
cream and bring to a boil, stirring constantly. Remove from heat and stir in vanilla. Store at
4°C. Heat before using in microwave or by placing container in a pan of hot water.

195

Appendix 1: Reagents

Kan Plates
5 g NaCl
10 g Bactotryptone
5 g yeast extract
15 g agar
DD H2O to 1 liter
Add 1 ml Kanamycin stock before pouring.
Kan Amp Plates
As for Kan plates, but add 0.4 ml amp stock.
Kamamycin Stock
Stock: 5 g Kanamycin (Sigma) in 100 ml H2O
Filter sterilize
add 1 ml / liter plates
1 M KCl
37.28 g KCl
Dissolve in 500 ml DDH2O
Autoclave.
5 M KOAc
49 g / 100ml H2O
L Broth
5 g NaCl
10 g bactotryptone
5 g yeast extract_____________
1 liter H2O
Autoclave.

10
20
10
__
2 liter

20
40
20
__
4 liter

Lambda Plates
Make up in 4 liter flasks. Autoclave 75 minutes with stir bar. Stir while cooling.
1l
2.5 l
NaCl
5g
12.5 gBacto-Tryptone
10 g
25 g
Yeast extract5g
12.5 g
Agar
11 g
28 g
MgSO4
1.2 g
3g
water to:1 liter
2.5 liter
Lysis Mixture

196

Appendix 1: Reagents

500 ml 250 mM EDTA, pH 8
100 ml 1 M Tris, pH 8
2 ml Triton X-100
Dilute to 2 liters with DD H2O.
N.B. Adjust the volume before mixing, due to sudsing.
Autoclave. This will be cloudy after autoclaving but will clear up as it cools.
20% maltose
400 g maltose
Dissolve in 2 liters DDH2O.
Autoclave.
Marker Stock
4 µg l Hind III
5 µg FX Hae III
TE to 400 µl
200 µl Ficoll-BPB with 25 mM EDTA
1 M MgCl2
101.6 g MgCl2 • 6 H2O
Dissolve in 500 ml DDH2O.
Autoclave.
0.1 M MgSO4
12g MgSO4
DDH2O to 1 liter
aliquots of 100 ml/bottle
Autoclave
Minimal Plates
for 500 ml: Autoclave 7.5g agar in 419 ml DDH2O with stir bar.
Add following sterile solutions:
50 ml 10xM9 phosphate
5 ml 10% NH4Cl
5.5 µl 0.1 M MgSO4
20 ml 10% glucose
1 ml 10 mg/ml thiamine HCl (add when ready to pour)
10X M9 Phosphate
30 g KH2PO4

197

Appendix 1: Reagents

113g Na2HPO4 • 7 H2O / to 1 liter H2O
Autoclave
M9 Mediumper liter:
100 ml 10X phosphate
10 ml 10% NH4Cl
11 ml 0.1 M MgSO4
40 ml 10% glucose
2 ml 10 mg/ml thiamine HCl
837 ml M9 CAA
5 M NaCl
146 g NaCl
to 500 ml with DD H2O
Autoclave.
(Note: heat to dissolve salt as concentration is near saturation.)
10% NH4Cl
200 grams NH4Cl
DD H2O to 2 liters
Autoclave.
4X Nick Translation Buffer
84 µl 1 M Tris pH 8
336 µl 1 M Tris pH 7.5
42 µl 1 M MgCl2
40 µg Pentex BSA
1498 µl H2O.
O/N Broth
20g Tryptone
10g NaCl
Yeast extract 10g
DD H2O to 1 liter
Autoclave.
add maltose to 0.1% before use if desired (0.5 ml 20% maltose/100 ml O/N).

198

Appendix 1: Reagents

10% SDS
100 g SDS (Biorad)DD H2O to 1 liter
DO NOT AUTOCLAVE
20X SSC
350.6 g NaCl
176.5 g NaCitrate
Dissolve in 2 liters DD H2O.
Autoclave.
20X SSCP
800 ml 1M sodium phosphate pH 6.8
176.4g NaCitrate
280.5g NaCl
to 2 liters with DD H2O.
Autoclave.
Sinsheimer's Medium
1.5 ml 5 M NaCl
9.0 ml 10% NH4Cl
36 ml 10% glucose
20 ml 20% maltose
5 ml 1 M MgCl2
2 ml 50 mM CaCl2
840 ml Sinsheimer CAA
3 M sodium acetate
246.1 g sodium acetate (anhydrous)/1 liter H2O.
Autoclave1 M sodium phosphate pH 6.8
138g
268g

NaH2PO4 • H2O
Na2HPO4 • 7 H2OAdjust to 2 liters with DD H2O.
Autoclave.

199

Appendix 1: Reagents

Southern Blot Sol'ns
Denat.

0.5 M NaOH
1.5 M NaCl

Neut.

1 M Tris pH 7.4
3M NaCl

4 liters
200 ml 10N NaOH
350.6 g NaCl
484.4 g Tris pH 7.4
701.3 g NaCl
pH with >250 ml conc HCl

Stabs
Bactotryptone
NaCl
Difco agar
6g
Yeast extract
DDH2O to 1 liter

8g
5g
1g

Adjust pH to 7.2-7.4 with about 4 drops of 6 M NaOH per liter. Autoclave 5 min to dissolve
agar. Fill vials with 3 ml each. Autoclave. Leave upright.
25% Sucrose (w/v) in 50 mM Tris HCl, pH 7.5
500g sucrose
100 ml 1 M Tris HCl, pH 7.5
DD H2O to 2 liters
Autoclave.
20X TBE (Tris-borate, electrophoresis buffer)
432 g Tris base
220 g Boric acid (crystal dissolves easier than powder)40.8 g Na2 EDTA • 2 H2ODissolve in 2 liters DD H2O
Check: pH = 8.35 for a 1:20 dilution
Autoclave. (Use Bellco 1 liter bottles)

200

Appendix 1: Reagents

TCM
5 ml 1 M Tris HCl, pH 7.5 (10 mM Tris HCl, pH 7.5)
100 ml 50 mM CaCl2 (10 mM CaCl2)
5.0 ml 1 M MgCl2 (10 mM MgCl2)
Dilute to 500 ml with DD H2O.
Autoclave.
TE pH 8.0 (10 mM Tris; 1 mM EDTA)
10 ml 1 M Tris pH 8.0
4 ml 250 mM EDTA, pH 8.0
Adjust volume to 1 liter with DD H2O.
Autoclave.
TE/0.2 M NaCl
11.6 g NaCl
Dissolve in 1 liter TE.
Autoclave.
Tetracycline
5 g tetracycline (tetracycline HCl, pharmaceutical grade)
100 ml 50% Ethanol / 50% sterile DDH2O
Want 25 µg/ml
Add 0.25 ml/500 ml L-broth
Let agar stir a while after adding antibiotic.
TM10 with Gelatin
2.41 g MgSO4
20 ml 1 M Tris pH 7.5
20 ml 1% gelatin (or 0.2g dry gelatin. Heat to dissolve.)
DD H2O to 2 liters.
Autoclave.
TM10 - 10 mM MgSO4 and 10 mM Tris pH 7.5
2.41 g MgSO4
20 ml 1 M Tris pH 7.5
Make up to 2 liters with DD H2O. Autoclave.
Top Agar201

Appendix 1: Reagents

5 g NaCl
10 g bactotryptone
5 g yeast extract
1.2 g MgSO4
7 g Agar DD H2O to 1 liter.
This must be heated to almost boiling. Aliquot 50 ml per milk dilution bottle. Autoclave.
Top Agarose
5 g NaCl
10 g bactotryptone
5 g yeast extract
1.2 g MgSO4
1 liter DD H2O
7 g Agarose (low EEO)
This must be heated to almost boiling. Aliquot 50 ml per milk dilution bottle. Autoclave.
300 mM Tris Base
36.34 g Tris base
to1 liter with DD H2O
Autoclave.
1 M Tris pH 8
177.6 g Trizma HCl
106 g Trizma baseDissolve in 2 liters DD H2O.
Autoclave.
1 M Tris pH 7.5
254 g Trizma HCl
47.20 g Trizma baseDissolve in 2 liters DD H2O.
Autoclave.

202

Appendix 1: Reagents

50 mM Tris pH 8
50 mM EDTA pH 8
15% sucrose (w/v)
100 ml of 1 M Tris pH 8
400 ml of 250 mM EDTA
300 g sucrose
DD H2O to 2 liters.
Autoclave.
2X YT Broth
16g bactotryptone
10g yeast extract
10g NaCl
H2O to 1 liter
Autoclave.
1X YT AgarTop
Bottom
8g
8g
5
5
5
5
7
15
___
___
1 liter
1 liter
Autoclave.

Bactrotryptone
yeast extract
NaCl
Agar
H2O

Carrier DNA
10 g herring testis DNA (Sigma Type XIV)DDH2O to 2 liters
Stir until all DNA is no longer opaque. Autoclave. Freeze until use.
100X Denhardt's Solution
20g polyvinylpyrolidone
20g bovine serum albumin (Sigma Type V)
20g ficoll
10ml 0.25M EDTA
H2O to 1 liter203

Appendix 1: Reagents

DNase for Nick-Translation
10-4 mg/ml Worthington DNase (DPFF) in 50% glycerol, 1X NT buffer.
Ethidium bromide
500 mg ethidium bromide (Sigma)/100 ml DDH2O
Loening Buffer
4.36 g Tris base (36 mM)
4.14 g NaH2PO4 • H2O (30 mM)
0.336 g Na2EDTA (1 mM)
DDH2O to 1 liter
Nucleotide triphosphates
To 25 mg nucleotide triphosphate add 350µl sterile water. Bring to pH 7 by adding 1N NaOH
(20-50 µl), using pH paper to test after each addition. Make a 1/2000 dilution into a buffer of theappropriate pH and measure the optical density at the appropriate wavelength. The final
concentration should be about 100mM.
Nucleotide

pH ofdilution buffer

Wavelength

Extinction
coefficient

dATP
ATP

7
7

259 nm259

15.2 x103
15.4 x103

dCTP
CTP
dGTP
GTP

2 (0.01 M HCl)
2
7
7

280
280
253
253

13.1 x 103
12.8 x 103
13.7 x 103
13.7 x 103

dTTP

7

267

9.6 x 103

UTP

7

262

10.0 x103

NOTE: conc. = optical density/extinction coeff. x dilution factor. So for a 100 mM solution of
UTP, diluted 1/2000 in pH 7 buffer, the optical density at 262 nm will be 0.5: 10-1 M = (0.5 O.D.
units/10x103 M/O.D.) x (2x103).

204

Appendix 1: Reagents

Restriction Enzyme Buffers
See Maniatis
10X Reverse Transcription Buffer
for 5 ml:

2.5 ml 1M Tris HCl, pH8.3 (500mM)
0.6 ml 5 M NaCl (600mM)
0.3 ml 1M Mgcl2
12.5 ml 2 M KCl

Make 100µl aliquots. Add 1µl 1M DTT/ 50µl reaction.

205

INDEXβ-mercapto ethanol 147
bacteriophage lambda bibrary screening
169
bacteriophage lambda DNA 166
basic fuchsin 60
Benton and Davis Solutions 193
Benzamidine 147
biotin labeled probes 26
biotin-16-dUTP 45
Bloomington Stock Center 4
Bluescript vectors 110
Boehringer Genius Kit 43
boiled RNAse 98
boiling method 124
Bowling Green Stock Center 4
BrdU 163
bromodeoxyuridine 163
bromophenol blue 148
BU buffer 193
BUdR 163
Buffer A+B 159

0.1M HEMG 159
10% Glucose 195
10X BU Buffer 193
10X kinase buffer 102
20 mM DTT 194
25% Sucrose (w/v) in 50 mM Tris HCl,
pH 7.5 200
250 mM EDTA 194
35S 333b-indole-acrylic acid 145
4M ammonium sulfate 160
50 mM CaCl2 194
500X GRACE'S VITAMINS FORECHALIER'S D22 MEDIUM 21
5X oligo-labelling buffer 108
5X PE 111
Acetyl CoA 187
acrylamide 152
bis 146
acrylamide SDS gel 146
Acrylamide stock 193
adjuvant 113
adult flies 158, 185
adult head 74
affinity purification of sera 142
alkaline lysis 124
Aminopterin 117
Ammonium persulfate 146
Amp Plates 193
ampicillin 145, 193
antibodies 55
antibody 145, 151
antibody production 145, 149
Antibody Staining 53, 76
antigen 151
antisera 142
AP conjugate 154
aprotinin 147
APS 152
ascites 120
Autoradiography 39

CaCl2 194
CAM 187
carcinogen 77, 80
Carnoy's fix 163
Carrier DNA 203
Casamino Acids for Sinsheimer's 194
CAT assay 187
cDNA cloning 98
cDNA library 172
cDNAs 145
cell fusion 117
Chloramphenicol Stock - 45 mg/ml 194
chromosome walking 188
cinnabar 80
CJ236 137
cloning of hybridoma 120
Clusters 81
Co 58
cobalt intensification 63
cobalt sulfide 64
cobalt sulphide staining 59
coomassie 146
coomassie brilliant blue 148
cosmid 188
Crictical point dry 92
cryoprotection 76
cryostat 74, 76
CsCl 193

β-galactosidase activity staining 74
β-Galactosidase Activity Staining using
X-gal 65, 72
β-galactosidase detection 65, 72
β-Galactosidase for EM Histology 69
β-galactosidase fusion proteins 139

206

INDEXDAB 77
deep pseudopupil 191
Denhardt's Solution 203
devitellinization of embryos 65
Dialysis Bags 194
diaminobenzidine 56
digoxigenin 51
Digoxigenin-11-dUTP 45
digoxigenin-labelled DNA 45
digoxigenin-UTP 43
DNA isolation 185
DNA labelling 109
DNA prep. 166
DNA sequencing, 130
DNA shearing 97
DNA-binding 104
DNAaseI footprinting 104
DNase for Nick-Translation 204
double-stranded sequencing 138
DPX 56, 75, 77
DPX mounting medium 73
drosophila tissue culture 18
DTT 194

fixation 52
Fixation and embedding 87
Fly Egg Laying Medium 195
fly food 6
FM6 80footprinting 102, 104
footprinting extracts 157
freezing cells 116
Freezing Medium 195
frozen tissue sections 74
fusion 118
fusion protein 139, 142
fusion proteins 145
G-10 spin column 130
G418 selection 17
Gelatin 195
genomic DNA preps 97
genomic library 171
Giemsa 41
glacial acetic acid 148
glucose 195
glutaraldehyde 74
glutathione 149
Glutathione-agarose beads 150
glycerol 148
glycerol mounting 75, 77
GST fusion proteins 149
guanidine hydrochloride 99

E. coli HB101 145
E. coli Trp E 145
ECHALIER'S D-22 MEDIUM FOR Kc
CELLS (HIGH PO4) 20
EDTA 147, 194
Egg Laying Medium 195
electro-transformation 161
electroelution 155
electrophoresis buffer 200
electroporation 161
ELISA 118
elution 155
Elution buffer 130
embryos 43, 52, 65
EMS 80
EMS mutagenesis 80
endcloning 188
ethane methyl sulfonate 80
Ethanol 148
Ethidium bromide 204
eye imaginal discs 55
false positives 81
FCS 114
Ficoll/Bromphenol Blue Loading Buffer
194
Film 192

207

Hanahan cells 101
HB Buffer 98
HB-4 rotor 146
HC110 94
head extracts 157
HEMG 159
high efficiency competent cells 101
Hoechst 75, 77
Hoechst 33258 54
homogenizing buffer 159
Hot Fudge Sauce 195
HRP 154
HRP-conjugated goat-anti-mouse IgG 77
hybridisation 26, 169, 175
hybridisation of oligonucleotides 176
hybridoma 151
imaginal disc 50
imaginal discs 72
Immunization 141
immunohistochemical staining 72

INDEX
immunostaining 52
in situ hybridization 26, 33
in situ hybridization with nonradioactive probe 43
ISCO apparatus 155
isolation of Drosophila DNA 185
isolation of Drosophila RNA 186
isopropyl gallate, 54

microscopy 87
Mid-America Drosophila Stock Center 5
milk 153
mini-prep 98, 123
Minimal Plates 197
monoclonal antibody 114
Mosaicism 81
mutagen 80
mutagenesis 80
Myeloma 114, 117

J774A.1 114
Kamamycin Stock 196
Kan Amp Plates 196
Kan Plates 196
Kc cell line. 18
KCl 196kinase buffer 102
KOAc 196

NaCl 147, 148, 198
NH4Cl 198
Ni 58
Nick Translation Buffer 198
non-radioactive in situ hybridization 50
non-radioactive in situs 33
Northern blots 110
Northerns 98
nuclease S1 133
Nucleotide triphosphates 204

L Broth 196
lacto-aceto orcein stain 23
lacZ fusion proteins 142
lambda agarose plates 166
lambda DNA 166
Lambda Plates 196
lambdasorb 165
Lämmli buffer 146, 148
leupeptin 147
LGT agarose 108
library screening 169
lifts 169, 173
Lithium 187
Loading Buffer 194
local mutagenesis 83
localization of RNA 43
Loening Buffer 204
Lysis Mixture 197
lysozyme 145

O.C.T. Tissue Tek 76
O.C.T. Tissue Tek compound 74
O/N Broth 198
Oak Ridge tube 97
oligo-labelled DNA probes 108
oligonucleotide 132
oligonucleotide primer 106
oligonucleotides 130
Optical Grade CsCl 193
P-element excision 83
P-element transformant 90
P[lacZ] lines 100
paraformaldehyde 33
Paraplast Plus 34
pATH 1 145
pATH 10 145
pATH 11 145
pATH plasmids 145
PBSG 76
PCM 115
PCR-labeled Single-stranded Probe 44
PEG 129
PEG 50% 117
PEM 55
PEM-formaldehyde 52
pepstatin 147
peripodial membrane 55
permeabilisation 52

M13 phage 128
M13 template DNA 133
M3 MEDIUM - FOR SCHNEIDER
LINE-2 CELLS 22
M9 Medium 198
M9 Phosphate 198
M9CA 145
maltose 197
Marker Stock 197
methanesulfonic acid ethyl ester 80
MgCl2 197
MgSO4 197

208

INDEX
peroxidase/DAB staining 73
PGB 148
pGEX 149
phage 128, 166
Phenylmethane-sulfonyl, in propanol
147
photomicrography 94
photoreceptor 191
pHSNEO 17
plaque purification 169
plaque purifying 181
plaque-purification 128
plasmid preps 122, 124, 126
plasmid rescue 100
Plate lysate 166
platinum 93
PLP 55PMSF 147
polyA+ RNA 98
polyclonal antibody 113
polytene chromosomes 23, 26
Ponceau's S stain 153
population cage 7
primer extension 107
primer labelling 106
probes for footprinting 102
protein gels 152
Proteinase K digestion 44
pseudopupil 191
pupal retina 63
purine cleavage markers 103
quarantine 4

Sectioning and Staining 88
seminar slides 192
Sequenase 138
serum 141
short-wave UV shadowing 130
signal intensification 73
Silica Gel TLC plate 132
silver-gold intensification 54
silver-gold method 58
single-stranded DNA 168
Sinsheimer's Medium 199
site-directed mutagenesis 133, 137
size standards 146
Slidemaker 192
slides 192
small scale plasmid DNA prep 124
small-scale DNA prep 166
snot 125
SOC medium 162
sodium acetate 199
sodium phosphate pH 6.8 199
somatic mosaic analysis 90
Southern Blot Sol'ns 200
SSC 199
SSCP 199
Stabs 200
Stock Center 4
STET 124
Stratagene transcription kit 36, 110
Subbing slides 35
sucrose 148, 200
sucrose infiltration 76
Sylgard 184 57, 64

radioactive in situs 33
Random hexamers 108
reagents 193
Restriction Enzyme Buffers 205
retinab-galactosidase activity 78
Reverse Transcription Buffer 205
RIBI 113
riboprobes 110
RNA isolation 99, 186
RNA prep 98
rosy 83
S.E.M. 92
saponin 54
scarlet 80
Schneider line 2 cell line 18
SDS 146, 199
SDS polyacrylamide 152

TBE Tris-borate, 200
TBS 153
TCM 201
TE pH 8.0 (10 mM Tris 201
TE/0.2 M NaCl 201
Technidol LC 94
tegosept 6
TEMED 146
Tetracycline 201
TG-2 137
thawing cells 116
tissue culture 18
TLC 132
TLC-purified oligonucleotides 132
Tm 179
TM10 - 10 mM MgSO4 and 10 mM Tris
pH 7.5 201

209

INDEX
TM10 with Gelatin 201
Top Agar 202
Top Agarose 202
TRANSFECTION 19, 128
transformation vectors 10
translational fusion 145
triphosphates 204
Tris 146
Tris 50mM,pH8.0, EDTA 50mM, pH8.0,
15% sucrose (w/v) 203
Tris Base 202
Tris-borate, 200
Tris pH 7.5 202
Tris pH 8 202
triton X-100 145, 147
Trp E fusion proteins 145
trpE fusions 143
TTBS 153

210

Vectastain 163
western blot 153
white 80, 83, 90
X ray-induced mitotic recombination 90
X-gal 74
X-gal crystals 66, 75
X-linked lethals 80
x-ray mutagenesis 86
Yeast-Glucose Plates 91
yellow, 80
YT Agar 203
YT Broth 203
Zwitergent 146, 147