Phenol-chloroform RNA isolation from Nematostella
- Collect embryos or small polyps by swirling them around in the dish,
pipetting off garbage, and pipetting embryos into 1.5 ml centrifuge tube,
minimizing water. Pipette off excess water.
- In the hood, add 1 mL Trizol to each tube of embryos. Swirl to dissolve;
vortex lightly if needed. Make sure all embryos are dissolved. If extracting
from small polyps, start with 0.5 mL Trizol, homogenize, then add another
0.5 mL Trizol.
Spin down heavy phase lock tubes, keeping centrifuge at 4°C. Transfer
the phenol containing dissolved embryos into the heavy phase lock tubes.
Add 200 μL chloroform to each tube and shake well for 15 seconds.
Incubate 10 minutes on ice.
Spin down at max speed for 15 minutes at 4°C.
Spin down new, empty phase lock tubes for 1 minute. Transfer the
aqueous phase to the new phase lock tubes.
Add 600 μL phenol-chloroform-isoamyl-alcohol to each tube and shake for
15 seconds.
Incubate on ice for 5 minutes.
Spin down at max speed 15 minutes at 4°C.
Transfer aqueous phase to new, clean 1.5 mL tube. Use a barrier tip and
be very careful the tube does not touch the bench.
Add 1 μl glycogen.
Add 500 μl isopropanol, shake, and incubate at room temperature 10-20
minutes.
Spin at max speed for 15 minutes at 4°C.
Remove liquid being careful to avoid pellet. Spin again for 10 seconds and
remove more liquid.
Add ~890 μl RNase-free 70% EtOH stored at -20°C. Vortex.
Remove liquid, avoiding pellet. Spin down briefly and remove liquid again.
Repeat (16) & (17).
Let pellet dry thoroughly and add 10μl RNase free water.
PART I: Select primers
• Ideal probes are the entire length of your gene. If the gene is >2 kb, the
probe can be a 1-2 kb fragment of the gene. Probes should be at least 1 kb.
• Using Geneious, select 200-250 bp at the beginning of your gene sequence.
Use “Design primers” and select forward primer. Primer should be:
o length: 20-27 bp; optimal 22 bp
o Tm: 57-63°C; optimal 60°C
o GC: 40-60%; optimal 50%
• Repeat for last 200-250 bp of gene, using “Design primers” and selecting
reverse primer.
• Blast primers against the species genome if possible to make sure they hit
only your target gene.
• Check for primer dimers using online tools.
PART II: Make template cDNA
Materials
• Trizol reagent
• chloroform
• phenol/chloroform/isoamyl alcohol
• glycogen
• isopropanol
• RNAse-free 70% EtOH
• Advantage RT-for-PCR cDNA synthesis kit
- Isolate RNA using Trizol reagent protocol.
- Collect embryos or small polyps by swirling them around in the dish,
pipetting off garbage, and pipetting embryos into 1.5 ml centrifuge tube,
minimizing water. Pipette off excess water.
- In the hood, add 1 mL Trizol to each tube of embryos. Swirl to dissolve;
vortex lightly if needed. Make sure all embryos are dissolved. If extracting
from small polyps, start with 0.5 mL Trizol, homogenize, then add another
0.5 mL Trizol.
Spin down heavy phase lock tubes, keeping centrifuge at 4°C. Transfer the
phenol containing dissolved embryos into the heavy phase lock tubes.
Add 200 μL chloroform to each tube and shake well for 15 seconds.
Incubate 10 minutes on ice.
Spin down at max speed for 15 minutes at 4°C.
Riboprobe synthesis for in situ hybridization
Spin down new, empty phase lock tubes for 1 minute. Transfer the
aqueous phase to the new phase lock tubes.
Add 600 μL phenol-chloroform-isoamyl-alcohol to each tube and shake for 15 seconds.
Incubate on ice for 5 minutes.
Spin down at max speed 15 minutes at 4°C.
Transfer aqueous phase to new, clean 1.5 mL tube. Use a barrier tip and be very careful the tube does not touch the bench.
Add 1 μl glycogen.
Add 500 μl isopropanol, shake, and incubate at room temperature 10-20 minutes.
Spin at max speed for 15 minutes at 4°C.
Remove liquid being careful to avoid pellet. Spin again for 10 seconds and
remove more liquid.
Add ~890 μl RNase-free 70% EtOH stored at -20°C. Vortex.
Remove liquid, avoiding pellet. Spin down briefly and remove liquid
again.
Repeat (16) & (17).
Let pellet dry thoroughly and add 10μl RNase free water.
Make cDNA using Advantage RT-for-qPCR kit. Follow published protocol
(below). Use 1 μg RNA.
PART III: Clone probe sequence into plasmid
Materials
Day 1
• Taq polymerase and 10x buffer
• dNTPs (10 mM)
• QiaQuick gel purification kit
• isopropanol
• pGEM-T plasmid ligation kit
Day 2
• DH5-alpha (competent bacterial cell line)
• LB liquid medium
• LB/ampicillin agarose plates
Day 3
• LB liquid medium with ampicillin
• Sp6 and T7 primers
Riboprobe synthesis for in situ hybridization
Day4
• 5 Prime Fast Plasmid kit
• 50% glycerol
-DAY 1-
- Amplify the probe sequence from cDNA using selected primers.
Reaction:
10x reaction buffer 5 μl
dNTPs (10 mM) 0.5 μl
Taq polymerase 0.5 μl
nuclease-free water 41 μl
forward primer (10 μM) 1.0 μl
reverse primer (10 μM) 1.0 μl
template cDNA 1.0 μl
TOTAL 50 μl
Amplify 40 cycles.
Run the PCR product on a 1% agarose gel. Extract the correct band using a
QiaQuick gel purification kit (Protocol below from D.K. Simmons)
Excise the DNA fragment from the agarose gel with a clean, sharp
scalpel.
Weigh the gel slice in a colorless tube. Add 400μl of Buffer QG to 1
volume of gel (100 mg gel ~ 100 μl). The maximum amount of gel slice
per spin column is 400 mg. (or just add 400 μl) For >2% agarose gels,
add 6 volumes Buffer QG.
- Incubate at 50°C for 10 min (or until the gel slice has completely
dissolved). Vortex the tube every 2–3 min during incubation to help
dissolve the gel.
- After the gel slice has dissolved completely, check that the color of the
mixture is yellow (similar to Buffer QG without dissolved agarose). If
the color of the mixture is orange or violet, add 10 μl 3 M sodium
acetate, pH 5.0, and mix. The color of the mixture will turn to
yellow.(~400ul)
Add 1 gel volume of isopropanol to the sample and mix by inverting.
(~133ul)
Place a MinElute spin column in a provided 2 ml collection
Apply sample to the MinElute column and centrifuge for 1 min or
Discard flow-through and place the MinElute column back into the
same collection tube. For sample volumes of more than 800 μl, simply
load and spin again.
- Add 500 μl Buffer QG to the MinElute column and centrifuge for 1 min.
Discard flow-through and place the MinElute column back into the
same collection tube.
- Add 750 μl Buffer PE to MinElute column and centrifuge for 1 min
Discard flow-through and place the MinElute column back into the
same collection tube.
Note: If the DNA will be used for salt-sensitive applications, such as
direct sequencing and blunt-ended ligation, let the column stand 2–5
min after addition of Buffer PE.
- Centrifuge the column in a 2 ml collection tube (provided) for 1 min.
Residual ethanol from Buffer PE will not be completely removed unless
the flow-through is discarded before this additional centrifugation.
- Place each MinElute column into a clean 1.5 ml microcentrifuge tube. To
elute DNA, add 10 μl water to the center of the MinElute membrane.
(Ensure that the elution buffer is dispensed directly onto the membrane
for complete elution of bound DNA.) Let the column stand for 1 min,
and then centrifuge the column for 1 min.
Ligate the fragment into the pGEM-T vector plasmid. (Protocol below from
D.K. Simmons)
Briefly centrifuge the pGEM®-T or pGEM®-T Easy Vector and Control
Insert DNA tubes to collect the contents at the bottom of the tubes.
Set up ligation reactions as described below.
Note: Use 0.5ml tubes known to have low DNA-binding capacity (e.g.,
VWR Cat.# 20170-310). Vortex the 2X Rapid Ligation Buffer
vigorously before each use.
Mix the reactions by pipetting. Incubate the reactions for 1 hour at room
temperature.
Alternatively, if the maximum number of transformants is required,
incubate the reactions overnight at 16°C.
Reaction Component Standard Reaction 1/2 Rxn ¾ Rxn
2X Rapid Ligation Buffer, T4 DNA
Ligase 5ul 2.5ul
3.75ul
pGEM®-T or pGEM®-T Easy Vector
(50ng) 1ul 0.5ul
0.75ul
PCR product Xul (3ul) X(1.5ul) X(2.25ul)
T4 DNA Ligase (3 Weiss units/μl) 1ul 0.5ul 0.75ul
nuclease-free water to a final volume
of 10ul 5ul
7.5ul
Note: if >1kb think about using 25ng of vector instead, due to the lowered ratio.
-DAY 2-
Transform the ligation product into competent bacteria (we use DH5-alpha),
and grow colonies overnight. (Protocol below from D.K. Simmons)
Add 2μl of each ligation to a 1.5ml microcentrifuge tube on ice
Remove tube(s) of frozen JM109 High Efficiency Competent Cells from
storage and place in an ice bath until just thawed (about 5 minutes). Mix
the cells by gently flicking the tube. Avoid excessive pipetting, as the
competent cells are extremely fragile.
Carefully transfer 50μl of cells into each tube prepared in Step 2 (use
100μl of cells for determination of transformation efficiency).
Gently flick the tubes to mix and place them on ice for 20 minutes.
Heat-shock the cells for 45–50 seconds in a water bath at exactly 42°C
(do not shake).
Immediately return the tubes to ice for 2 minutes.
Add 460μl room-temperature LB medium to the tubes containing cells
transformed with ligation reactions and 900μl to the tube containing
cells transformed with uncut plasmid
- Incubate for 1.5 hours at 37°C with shaking (~150rpm).
- Plate 100μl of each transformation culture onto LB/ampicillin plates. If a
higher number of colonies is desired, the cells may be pelleted by
centrifugation at 1,000 × g for 10 minutes, resuspended in 200μl of LB
medium, and 100μl plated on each of two plates.
- Incubate the plates overnight (16–24 hours) at 37°C. If 100μl is plated,
approximately 100 colonies per plate are routinely seen using competent
cells that are 1 × 108cfu/μg DNA. Use of ultra-high- efficiency
competent cells may result in a higher number of background colonies.
Longer incubations or storage of plates at 4°C (after 37°C overnight
incubation) may be used to facilitate blue color development.
-DAY 3-
- Perform colony PCR to determine which colonies contain PCR fragment.
Reaction:
10x reaction buffer 2.5 μl
dNTPs (10 mM) 0.25 μl
Taq polymerase 0.5 μl
nuclease-free water 19.75 μl
Sp6 primer (10 μM) 1.0 μl
T7 primer (10 μM) 1.0 μl
TOTAL 25 μl
Touch a pipette tip to a colony, then dip it into the PCR solution.
Amplify for 40 cycles.
Set up liquid cultures for colonies containing PCR fragment.
Pour 3 ml LB into a glass culture tube.
Touch a pipette tip to the chosen colony, then drop it into the LB in the
tube.
Incubate overnight with shaking.
-DAY 4-
Miniprep liquid cultures using 5 Prime Fast Plasmid kit. Simultaneously
make glycerol stocks of the lines.
Chill the Complete Lysis Solution on ice.
Pellet 1.5 ml of fresh bacterial culture at maximum speed (at least 12,000
x g or 13,000 rpm) for 1 minute in the provided 2 ml Culture Tube.
Remove medium by decanting, taking care not to disturb bacterial
pellet.
Add 400ul of ICE-COLD Complete Lysis Solution. The Complete Lysis
Solution MUST be ice-cold (0 –4°C) to obtain maximum yield.
Mix thoroughly by constant vortexing at the highest setting for a full 30
seconds. This step is critical for obtaining maximum yield.
Incubate the lysate at room temperature for 3 minutes. (add 125ul of
Isopropanol)
Transfer the lysate to a Spin Column Assembly by decanting or
pipetting.
Centrifuge the Spin Column Assembly for 30– 60 seconds at maximum
speed.
Add 400 ul of DILUTED Wash Buffer to the Spin Column Assembly.
Centrifuge the Spin Column Assembly for 30–60 seconds at maximum
speed.
Remove the Spin Column from the centrifuge and decant the filtrate
from the Waste Tube. Place the Spin Column back into the Waste Tube
and return it to the centrifuge.
Centrifuge at maximum speed for 1 minute to dry the Spin Column.
Transfer the Spin Column into a Collection Tube.
Add 50 ul of Elution Buffer directly to the center of the Spin Column
membrane and cap the Collection Tube over the Spin Column.
Centrifuge at maximum speed for 30 – 60 seconds.
Remove and discard the Spin Column.
The eluted DNA can be used immediately for downstream applications
or stored at -20°C.
Sequence miniprep results to ensure plasmid insertion is correct. Save
glycerol stocks of bacteria containing correct plasmid.