PLASTIC SECTION
Day 1
Tissue Fixation with 4% Paraformaldehyde
Make required 1X PBS (0.1 M Sodium Phosphate Buffer Solution, pH: 7.0) and pH up to
11.0 using NaOH. Heat to 60-70ºC and then add paraformaldehyde in the hood. Cool on
ice and pH back down to 7.0 with H2SO4.
*For 20 mL, add 290 uL 5 N NaOH to pH: 11, microwave for 20 sec, add 0.4 g
paraformaldehyde, put on ice, and then add 20 uL H2SO4 to pH: 7.0*
Tissue Types:
For siliques, slice siliques in half-sections and stick onto tape. Cut tape, gently roll to fit
into microcentrifuge tube and add the fixative solution made fresh previously. Make sure
that tissue is completely submerged in the fixative.
Day 2
Dehydration
Wash twice with 1X PBS. Dehydrate at 1 hr intervals (unless otherwise noted) with
ethanol in the following order:
 30% EtOH
 50% EtOH
 70% EtOH
 95% EtOH
 100% EtOH twice at 30 minutes and leave overnight
 100% EtOH the next day before going onto the Embedding step
 Addition dehydration with 100% is recommended to remove excess chlorophyll
Day 3
Infiltration (Embedding can be done on the same day)
1. Make Infiltration Solution by adding 1.25 g benzoyl peroxide, plasticized catalyst
(Polysciences, Inc. Cat #02618) to100 mL JB-4 Solution A (Electron Microscopy
Sciences Cat #14270-01). Exact measuring of benzoyl peroxide is crucial because
it controls rate of polymerization.
2. Stir with stir bar until catalyst is completely dissolved (about 15 min).
3. Infiltrate by doing ascending grades of the Infiltration Solution mixture with
ethanol in the following concentrations for 30 min incubation between each step:
50% ethanol + 50% Infiltration Solution
25% ethanol + 75% Infiltration Solution
10% ethanol + 90% Infiltration Solution
3x wash with the Infiltration Solution
It is recommended to have the tubes on a slow rotator/shaker during incubation.
Also, the volume of the Infiltration Solution should be 8-10 times that of the
tissue specimens. For specimens that fit in a 1.5 mL microcentrifuge (e.g. cut-up
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siliques, stems, roots, etc.), 1 mL Infiltration Solution is sufficient. When
infiltration is complete, samples should be translucent and somewhat
hard/plasticized. Infiltration Solution can be stored in 4ºC for about a month for
future infiltration.
Embedding
If embedding is not done on the same day as infiltration, make a fresh 25 mL batch of
Infiltration Solution using 25 mL JB-4 Solution A + 0.312 g benzoyl peroxide,
plasticized. Remember to stir with stir bar.
1. Remove Infiltration Solution from the tissue specimen tubes.
2. Using forcep, carefully arrange specimens into mold pit.
3. To the 25 mL Infiltration solution, accurately pipette 1.0 mL JB-4 Solution B.
4. Do a quick stir with the stir bar and immediately add 1.5 mL of the mixture to the
molds. Add carefully as to not disturb specimens.
5. Gently push in block holders (EMS Cat # 70175-50). This step can be done before
achieving the mixture (Step 3) but bubbles may float towards the block base
(which does not affect the sectioning pit).
6. Arrange mold(s) into vacuum apparatus (under 15 psi) and leave overnight at in
recommended 4°C (or room temperature, but block may not be as solid).
Sectioning
The key to getting nice, thin 1 um sections is to avoid getting the block moist or wet. For
smaller pointed blocks, successive sections can form nice ribbons. However, for wider
molds, it is recommended that you use a razor blade to cut a flat surface. This flat surface
must be parallel to the glass knife’s edge for perfect sections, otherwise you can crack or
even break the block. For wider blocks, I DO NOT fill the boat with water as this tends to
leak through the back of the knife, making the block moist and the sections crackled.
Instead, I add very little water to the base of the boat, brush up the water to the knife’s
edge which will allow the sections to stick to the knife’s edge after each microtome turn.
I then use a sharp forcep to gently transfer each individual section to a slide that is placed
on a slide warmer, with the slides filled with drops of water. The heated water will create
enough entropy to unfurl the sections so they are flat with the slide.
Staining
The slides were stained for only 5 minutes in 0.2% Toluidine Blue and de-stained
overnight (longer de-staining is recommended, sometimes over the weekend). Change the
water at least twice using distilled water. The slides are then allowed to dry at room
temperature and visualized under a microscope.
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X-Gal Overlay for Y2H
To make 1-2 plates, 0.125 g agarose is dissolved in 25 mL Z-buffer (60 mM
Na2HPO4·2H2O, 40 mM NaH2PO4·H2O, 10 mM KCl, 1 mM MgSO4·7H2O, pH: 7.0 and
autoclaved) by microwaving. Solution is then cooled down to 50ºC and 0.5 mL of 10%
SDS and X-Gal in DMF (final concentration of 2 mg/mL) are added. Solution is poured
over -TLHA and incubated for approximated 30 minutes to an hour at 37ºC. Take
pictures immediately after that (do NOT wait until the next day because bacteria will start
to grow on the plate).
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Embryo Clearing With Hoyer’s Solution
From Liu C. M. and Meinke D. W. 1998. The titan mutants of Arabidopsis are distrupted
in mitosis and cell cycle control during development. Plant J. 16:21-31.
Day 1
Making Hoyer’s Solution (60 mL)
 7.5 g gum Arabic (Sigma, G9752)
 100 g chloral hydrate (Sigma C8383, controlled substance)
 5 mL glycerol (Sigma G6279)
 60 mL distilled water
Dissolve all the above components overnight. Solution can be stored at room temperature
indefinitely.
Day 2
1. Place 2-3 drops of the Hoyer’s solution onto a slide
2. Open siliques carefully with a pair of sharp forceps and transfer ovules to the
Hoyer’s on the slide. Make sure not to crush the ovules and the ovules should be
easily transfer since they will readily stick to the forcep
3. Gently place a cover slip over the samples
4. Incubate the slides for about 30 minutes
5. Observe the samples using a microscope equipped with Nomarski optics in
Heven’s lab
Single Embryo Genotyping Using FTA Cards
1. Ovules are individually transferred to FTA cards using a forcep, being careful not
to crush the ovules which can contaminate the forcep
2. Once all are transferred onto the card, a strip of parafilm is cut to size, placed on
top of samples and quickly banged with the end of a 50 mL tube (or any hard and
big tool is sufficient). Be careful not to crush and tear the parafilm
3. The samples are allowed to dry for an hour and punched out using the red disc
punching tool provided by the FTA kit
4. The discs are transferred to individual PCR tubes and washed twice with 20 uL
FTA Purification Reagent at 2 min per wash and while shaking
5. The discs are washed twice with 100 uL TE buffer at 2 min per wash
6. The discs are now ready to be used in lieu of DNA (although the company
suggest air drying, air drying is not recommended as it may give unreliable
results)
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Pathogen Infection Assay:
Testing for Basal Resistance using Pseudomonas syringae DC3000
From Xiao Lab
1. Grow Arabidopsis for 4-6 weeks under short day conditions (8 hours light). Plants
should have nice, green, and large rosette leaves and not bolted.
2. Inoculate Pseudomonas syringae ES4326 DC3000 wild-type bacteria into 80 mL
of LB broth that has been supplemented with a final concentration of 100 ug/mL
streptomycin (that has been filtered sterilized). Newly grown bacteria taken from
the -80°C, plated, and then inoculated is the best option as you will have growth
the next day. Older bacteria on plates that have been sitting in the fridge for over a
month will grow very slow, sometimes taking approximate 2-3 days.
3. Grow flask of bacteria in 30°C shaker overnight
4. Spin down the bacteria at 4000 RPM for 5’ and wash twice with 5 mL of 10 mM
MgCl2, making sure to re-suspend in by vortexing and centrifuge before and after
each wash
5. Re-suspend in MgCl2 and dilute the sample to achieve an OD600=0.0002
6. Mark the leaves to be infected using a black marker and putting a dot at where the
syringe will be placed
7. Using a 1 mL needle-less syringe, suck up 100 uL of the dilution and infiltrate the
abaxial leaf surface, being careful not to break/tear the leaf. You should be able to
see the liquid as it infiltrates the leaf from the top, casting a shadow. Wipe off
excess bacteria solution from the surface.
8. Put the plants back in the short-day growth chamber for an additional 3 days
9. To determine bacterial counts with the best statistics, I use only one leaf per
infected plant per genotype. Using the back side of a 200 uL pipette tip (which is
approximate 1 cm in diameter), I drill the leaf with the back side, and using a
forcep, place the rounded sample into a sterile tube
10. The leaf discs are first macerated using a plastic pestle for 20 seconds before 200
uL of 10 mM MgCl2 is added, and then macerated again using the same pestle for
an additional 10 seconds
11. Make serial dilutions of all the samples. Be sure not to get the samples mixed up
by labeling them correctly –I have done as many as 30 samples successfully. The
1st dilution is a 50X dilution, which is 20 uL of the sample diluted in 980 uL of
dH2O. The 2nd dilution is a 100X dilution, which is 10 uL of the previous dilution
in 990 uL of dH2O. Make sure to vortex each sample before making the
dilutions immediately before to prevent bacterial sedimentation, which can
skew the results
12. The 2nd dilution is already diluted 5,000X. From that 2nd dilution, pipette out 100
uL and plate onto LB plates supplemented with a final concentration of 100
ug/mL of streptomycin. Because only 1/10th of the dilution was used, the final
dilution factor is 50,000X.
13. Incubate plates in 30°C
14. In about 2-3 days after incubation, colonies should have already been formed.
Count the number of colonies and multiply the total for each sample plate by
50,000 to get the CFU (colony forming units)
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Amplifluor SNP-typing
Adapted from Chunxin (Black) Wang and Chemicon’s Amplifluor SNPs Genotyping
System for Assay Development [Protocol]
1. Order kit (S7907) from Chemicon
2. Order desired primers using the Amplifluor AssayArchitect website at
www.assayarchitect.com
3.
4. Extract DNA from tissue and dilute samples to 25 ng/uL with dH2O, otherwise
the concentration will be too much and not be able to distinguish between wildtype/mutant alleles
5. Make at least two cocktail preps using the following (per reaction), one for the
kit’s control and the other for the gene-specific/allelic discrimination unknowns
 4.6 uL dH2O
 0.5 uL of 5 uM FAM primer
 0.5 uL of 5 uM JOE primer
 0.5 uL of Control Primer Mix* (*or SNP Allele Specific Primer Mix)
 1.0 uL Reaction Mix S+
 0.8 uL 2.5 mM dNTPs
 0.1 uL Invitrogen Platinum Taq (Cat# 11304-011)
 2 uL of 25 ng/uL DNA* (*or dH2O for Non-Target Controls –NTC)
*Be sure to have 3 NTCs (2 uL water) per primer set*
6. PCR using the following program:
a) Pre-heat block to 96°C
b) Denature for 4 min at 96°C
c) Amplification #1: 20 cycles
a. 15 sec @96°C
b. 5 sec @57°C
c. 10 sec @72°C
d) Amplification #2: 22 cycles
a. 15 sec @96°C
b. 20 sec @55°C
c. 40 sec @72°C
e) Extension for 3 min @72°C
f) Hold at infinity at 20°C
5. Use BioRad Plate reader in Coleman’s lab. They have the manual for “End Point
Analysis” for discriminating between the different alleles. Follow along with the
manual (page 68 of the manual). The samples are read under “Data Analysis
Module” since you will do a post-PCR read
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Sterilizing Arabidopsis thaliana Seeds
1. Transfer desired quantity of seeds to sterile centrifuge tubes.
2. Add 200 µL of 70% ethanol.
3. Add 600 µL of dH2O to each tube (the seeds should not be exposed to 70%
ethanol for more than 30 seconds; thus, it is recommended that no more than 10
tubes of seeds be done at any given time).
4. Pipette out 600 µL of the supernatant, being careful not to remove too many
seeds. Make sure you change the tip after each tube to prevent seed
contamination.
5. Repeat steps 3-4 three times to further dilute the ethanol.
6. Add 200 µL of 0.6% sodium hypochlorite (aka bleach, which comes standard at
6%, so dilute it 10X). At this point, each tube should have 400 µL.
7. Add 600 µL of dH2O to each tube (the seeds should not be exposed to 0.6%
sodium hypochlorite for more than 30 seconds).
8. Pipette out 600 µL of the supernatant, being careful not to remove too many
seeds. Make sure you change the tip after each tube to prevent seed
contamination.
9. Repeat steps 7-8 three times to further dilute the sodium hypochlorite, but after
the final wash, DO NOT REMOVE the supernatant.
10. If transferring to MS plates, pipette the seeds and as much of the supernatant as
the media will allow for even distribution. If transferring to soil, do a quick vortex
and immediately pipette contents directly onto soil, making sure to move the
pipette to evenly space. Be gentle for this final step to prevent seed damage and
clogging of the pipette tips.
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