Jan 22, 2016
FISH Protocol for FISH & FLOW
- Matthew Sullivan1
- 1Matthew Sullivan Lab, University of Arizona, Ohio State University
- VERVE Net
- Sullivan Lab
Protocol Citation: Matthew Sullivan 2016. FISH Protocol for FISH & FLOW. protocols.io https://dx.doi.org/10.17504/protocols.io.dfe3jd
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
Created: July 17, 2015
Last Modified: March 26, 2025
Protocol Integer ID: 1222
Keywords: fish protocol for fish, fish protocol, fish, protocol, flow
Guidelines
Sample Collection
Biosphere 2 Samples
Materials
• Acid-Washed Glass Bottles (Enough for samples + 1 for collecting ocean water)
• 16% Formaldehyde
• 200 Proof Ethanol
• Ice & Cooler for transport (or 4°C equivalent)
• Serological Pipettes (25mL)
• Electronic Pipettor
Protocol
1. Submerge inverted capped acid-washed glass below the surface of the B2 ocean.
2. Remove cap while submerged and allow water to fill glass bottle. Swirl and empty.
3. Repeat Steps 1 & 2, but this time keep the seawater. This is your sample
4. Call B2 Energy Center to obtain B2 ocean temperature & salinity at time of sampling.
5. Add 70mL of 16% Formaldehyde to a new acid-washed glass bottle.
6. Add your sample to the bottle to q. to 1 liter.
7. Again repeat Steps 1 & 2 and keep the seawater.
8. Add 500mL of 200 Proof Ethanol to a new acid-washed glass bottle.
9. Add your sample to the bottle to q. to 1 liter.
10. Right before you leave the B2 Ocean, repeat Steps 1 & 2 and keep the seawater.
11. Add 1 liter of seawater to a new acid-washed glass.
Filtration Process
Materials
• 3-place filtration manifold (stainless steel or polyurethane)
• 3 glass microanalysis filter holder assemblies 47mm (tower, clamp, and base)
• 1 glass microanalysis filter holder assembly 90mm (tower, clamp, and base)
• 90mm GF/D Filter
• 47mm cellulose nitrate 0.45µm supor filter
• 47mm 0.2µm GTTP Isopore membrane filters
• 2 2 liter glass flasks with arm
• Tubing
• Vacuum pump
Protocol
1. Attach a 2 liter glass flask with arm to the vacuum pump using tubing.
2. Set up the 90mm filter holder assembly and add the 90mm GF/D filter.
3. Add your sample to the tower and turn on the vacuum to begin filtration. Make sure the vacuum pressure does not exceed 5 in Hg. The filtered product is called the pre-filtered product.
4. Set up the 47mm filter holder assembly with the flask and add the 0.2um GTTP Isopore membrane filters shiny-side up.
5. Filter the 15mL pre-filtered product per membrane filter (x3). Filter 100mL pre-filtered product per membrane filter (x9).
6. Use tweezers to remove membrane filters from the filter holder assembly and put on blotting paper in the dark shiny side up.
7. Once the membrane filter is dry, store in a petri dish.
CARD FISH
Reconstituting FISH probes
Materials
• Lyophilized Probe
• PCR water
• Ice & Ice Bucket
• Nanodrop System
Protocol
1. Add 100µL PCR water to lyophilized probe.
2. Incubate on ice for > 2 hours.
3. Finger-flick a few times and shake down material to bottom of tube. (NOTE: you do not want to vortex because HRP enzyme attached to the probe may detach)
4. Use 1.5µL of reconstituted probe and check the UV at 260 and 404 wavelengths on the Nanodrop.
5. Add PCR water depending on Nanodrop results.
Embedding Samples
***NOTE: For microscopy samples only!
Materials
• Low Gelling Point Agarose
• Parafilm
• Glass Slide
• 80-96% Ethanol
• Tweezers (Non-fine)
• 2 Large-sized Petri Dishes
• Sample filtered on 47mm 0.2µm GTTP Isopore membrane filters
Protocol
1. Pre-warm a petri-dish.
2. Boil 0.1% low gelling point agarose and pour into the pre-warmed petri-dish.
3. Let agarose cool down to 35-40°C.
4. Cover glass slides with layer of parafilm so that there is an even surface.
5. Using sterile tweezers, dip filter with both sides in the agarose and place it face-down (ie. shiny-side/bacteria-side down!) onto the parafilm-covered slide.
6. Let dry at room temperature.
7. Remove filter from slide surface by soaking parafilm covered slide in petri-dish filled with 80-96% Ethanol.
8. Air-dry filter on a kimwipe.
Pre-Treatment
Materials
• Lysozyme Solution (10mg/mL)
• Recipe (10mL):
• 1mL 0.5M EDTA pH 8.0
• 3mL 1M Tris-HCl pH 8.0
• 6mL MQ water (autoclaved)
• 100mg Lysozyme (Fisher Cat#BP535-5)
• 0.01M HCl
• Water Bath
• Tweezers (Non-fine)
• Sterile Scalpel
• Blotting paper
• Pencil
• 2 Medium-sized Petri Dishes
• Sample filtered on 47mm 0.2µm GTTP Isopore membrane filters
Protocol
1. Turn on water bath and pre-warm to 37°C and sterilize tweezers with 70% EtOH.
2. While the water bath is warming up, take out a piece of blotting paper. Put your sample filter shiny-side up on the blotting paper. Cut your sample filters in half using sterile scalpel and sterile tweezers to stabilize. Mark the filter piece using a pencil.
3. Make fresh lysozyme solution and pour it into a petri dish.
4. To permeabilize your sample, submerge your samples in the lysozyme solution. Wrap parafilm around the edge of the petri-dish.
5. Put the petri-dish into the 37°C water bath for 1 hour.
6. While waiting for your incubation, pour 0.01M HCl in a new petri-dish.
7. Once your incubation is over, put your sample filters in a strainer and rinse off any residual lysozyme solution with distilled water.
8. To inactivate of endogenous peroxidases, submerge your sample in the 0.01M HCl and incubate for 15 minutes.
9. Again, put your sample filters in a strainer and rinse off any residual lysozyme solution with distilled water.
10. Air dry on a kimwipe.
Hybridization
Materials
• Probe
• Hybridization Chamber Mix
• Recipe (2mL)
• % Formamide
• q. to 2mL with MQ water
• Hybridization Buffer
• Recipe (refer to Elke’s recipe)
• Hybridization Ovens
• Tweezers (Non-fine)
• Sterile Scalpel
• Blotting paper
• Pencil
• Sample filtered on 47mm 0.2µm GTTP Isopore membrane filters
• 50mL falcon tubes
• Styrofoam 50mL falcon tube holder
• Cardboard
• Glass slides
• Kimwipes
• Parafilm
Protocol
1. Pre-heat hybridization ovens to 46°C and 48°C.
2. Create a hybridization humidity chamber setup (refer to Fig.1).
3. Based on the probe you plan to use for hybridization, determine the % formamide to use.
4. Create the individual hybridization humidity chambers (Fig. 2) by inserting a kimwipe soaked in the correct 2mL % formamide-water mix (Hybridization Chamber Mix). Cover glass slides evenly with parafilm. Do NOT add slide to individual chamber yet.
a. eg. If probe needs 35% formamide, create hybridization chamber mix that is 35% formamide (700µL formamide, 1300µL MQ water)
5. Take out a piece of blotting paper. Put your sample filter shiny-side up on the blotting paper. Cut your sample already halved filters into eighths using sterile scalpel and sterile tweezers to stabilize. Number the filter pieces using a pencil with a “#” followed by “•”.
a. NOTE: The “•” indicates the orientation of the filter, once the “shine” is no longer obvious
6. Mix hybridization buffer with probe working solution [50ng DNA µl-1] in a 300:1 ratio.
7. Dip each filter completely into the Hybridization mix and place filters face-up on the parafilm covered glass slide. Spread the rest of the solution evenly onto the filters.
8. Put glass slide horizontally into individual hybridization humidity chamber with corresponding % formamide to probe.
9. Incubate at 46°C overnight.
Washing
*** Prepare wash buffer at same time as setting up hybridization
Materials
• Washing Buffer
• Recipe (refer to Elke’s recipe)
• 1x PBS
• H2O2
• Amplification Buffer
• Recipe (refer to Elke’s recipe)
• Alexa 488
• EtOH
• Ice
• Tweezers (Non-fine)
• Large-sized Petri Dishes
• 50mL Falcon Tubes
• Glass slides
• Kimwipes
• Parafilm
Protocol
Prior Day
1. Make 50mL Washing Buffer in 50mL falcon tube with corresponding % formamide to probe.
2. Warm at 48°C overnight.
Day Of
3. Cover glass slides evenly with parafilm. Set aside until later use.
4. Remove filters from individual humidity chambers and put in corresponding % formamide pre-warmed Washing Buffer. Incubate at 48°C for 10 minutes.
5. Pour 1x PBS in a Large-sized petri dish.
6. Transfer filters to 1x PBS and incubate for 15 minutes at room temperature.
7. Get Ice and put H2O2, Amplification Buffer, Alexa 488 in covered ice bucket.
8. Mix 1µL H2O2 with 199µL 1x PBS.
9. Create the Substrate mix in a 1000 Amplification Buffer: 10 diluted H2O2 : 3.3 Alexa 488 ratio. Keep in covered ice bucket until ready to use.
a. Eg. Substrate Mix: 1000µL Amplification Buffer, 10µL diluted H2O2, 3.3µL Alexa 488.
10. Take filters out of 1x PBS and quickly wipe off excess 1x PBS on kimwipe. Make sure NOT to touch to cell-side to the kimwipe.
11. Dip filters into the Substrate mix and put on parafilm covered slide. Spread the rest of the mix evenly onto the filters.
12. Put slides into large-sized petri dishes and seal petri dish with parafilm and put in 46°C for 45 minutes in the dark.
13. Dry filters on kimwipe and put in 1x PBS for 10 minutes at room temperature in the dark.
14. Pour MQ water into a Large-sized petri dish and pour 96% EtOH into a different Large-sized petri dish.
15. Transfer filters to MQ water and cover in dark for 1 min at room temperature.
16. Transfer filters to 96% EtOH and cover in dark for 1 min at room temperature.
17. Put on kimwipe on a paper towel and let dry covered in the dark.
18. Immediately begin resuspension step! NOTE: If for microscopy, you can store filters at -20°C.
Resuspension of Cells from Filters
Materials
• 150mM NaCl
• 10% Tween 80 Ampules
• 0.2µm syringe filter
• Luer-lok syringe
• Large-sized petri dish
• 2mL centrifuge tubes
• Tape
• Incubator
• Vortexers
Protocol
1. Pre-heat incubator to 37°C.
2. Pour 1x PBS in a Large-sized petri dish. Transfer hybridized filters to 1x PBS and incubate for 15 minutes in the dark at room temperature.
3. 0.2µm filter sterilize the 150mM NaCl.
4. Make Resuspension Buffer (30mL 150mM NaCl; 160µL 10% Tween 80)
5. Put 1.5mL Resuspension Buffer per 2mL centrifuge tube and put filter in centrifuge tube.
6. Incubate centrifuge tube for 30 minutes at 37°C shaking horizontally.
7. Tape Tubes to vortexer horizontally (6 tubes per for vortexer). Shake in dark for 15 minutes at 2500 rpm.
8. Remove filter (BUT keep filter just in case). The cells should now be in the buffer!!
.
Troubleshooting
Before start
Prepare wash buffer at same time as setting up hybridization.
Sample Collection
Submerge inverted capped acid-washed glass below the surface of the B2 ocean.
Remove cap while submerged and allow water to fill glass bottle.
Swirl and empty.
Repeat Steps 1-3, but this time keep the seawater.
Note
This is your sample.
Call B2 Energy Center to obtain B2 ocean temperature & salinity at time of sampling.
Add 70mL of 16% Formaldehyde to a new acid-washed glass bottle.
Add your sample to the bottle to q. to 1 liter.
Again repeat Steps 1-3 and keep the seawater.
Add 500mL of 200 Proof Ethanol to a new acid-washed glass bottle.
Add your sample to the bottle to q. to 1 liter.
Right before you leave the B2 Ocean, repeat Steps 1-3 and keep the seawater.
Add 1 liter of seawater to a new acid-washed glass.
Filtration Process
Attach a 2 liter glass flask with arm to the vacuum pump using tubing.
Set up the 90mm filter holder assembly and add the 90mm GF/D filter.
Add your sample to the tower and turn on the vacuum to begin filtration.
Note
The filtered product is called the pre-filtered product.
Note
Make sure the vacuum pressure does not exceed 5 in Hg.
Note
Make sure the vacuum pressure does not exceed 5 in Hg.
Set up the 47mm filter holder assembly with the flask and add the 0.2um GTTP Isopore membrane filters shiny-side up.
Filter the 15mL pre-filtered product per membrane filter (x3).
Filter 100mL pre-filtered product per membrane filter (x9).
Use tweezers to remove membrane filters from the filter holder assembly and put on blotting paper in the dark shiny side up.
Once the membrane filter is dry, store in a petri dish.
Reconstituting FISH probes
Add 100µL PCR water to lyophilized probe.
Incubate on ice for > 2 hours.
Finger-flick a few times and shake down material to bottom of tube.
Note
NOTE: you do not want to vortex because HRP enzyme attached to the probe may detach.
Use 1.5µL of reconstituted probe and check the UV at 260 and 404 wavelengths on the Nanodrop.
Add PCR water depending on Nanodrop results.
Embedding Samples (For microscopy samples only)
Pre-warm a petri-dish.
Boil 0.1% low gelling point agarose and pour into the pre-warmed petri-dish.
Let agarose cool down to 35-40°C.
Cover glass slides with layer of parafilm so that there is an even surface.
Using sterile tweezers, dip filter with both sides in the agarose and place it face-down (ie. shiny-side/bacteria-side down!) onto the parafilm-covered slide.
Let dry at room temperature.
Remove filter from slide surface by soaking parafilm covered slide in petri-dish filled with 80-96% Ethanol.
Air-dry filter on a kimwipe.
Pre-Treatment
Turn on water bath and pre-warm to 37°C and sterilize tweezers with 70% EtOH.
While the water bath is warming up, take out a piece of blotting paper.
Put your sample filter shiny-side up on the blotting paper.
Cut your sample filters in half using sterile scalpel and sterile tweezers to stabilize.
Mark the filter piece using a pencil.
Make fresh lysozyme solution and pour it into a petri dish.
To permeabilize your sample, submerge your samples in the lysozyme solution.
Wrap parafilm around the edge of the petri-dish.
Put the petri-dish into the 37°C water bath for 1 hour.
01:00:00
While waiting for your incubation, pour 0.01M HCl in a new petri-dish.
Once your incubation is over, put your sample filters in a strainer and rinse off any residual lysozyme solution with distilled water.
To inactivate of endogenous peroxidases, submerge your sample in the 0.01M HCl and incubate for 15 minutes.
00:15:00
Again, put your sample filters in a strainer and rinse off any residual lysozyme solution with distilled water.
Air dry on a kimwipe.
Hybridization
Pre-heat hybridization ovens to 46°C and 48°C.
Create a hybridization humidity chamber setup.
Note
Refer to Fig.1 in guidelines.
Based on the probe you plan to use for hybridization, determine the % formamide to use.
Create the individual hybridization humidity chambers (Fig. 2 in guidelines) by inserting a kimwipe soaked in the correct 2mL % formamide-water mix (Hybridization Chamber Mix).
Note
eg. If probe needs 35% formamide, create hybridization chamber mix that is 35% formamide (700µL formamide, 1300µL MQ water).
Cover glass slides evenly with parafilm. Do NOT add slide to individual chamber yet.
Take out a piece of blotting paper.
Put your sample filter shiny-side up on the blotting paper.
Cut your sample already halved filters into eighths using sterile scalpel and sterile tweezers to stabilize.
Number the filter pieces using a pencil with a “#” followed by “•”.
Note
NOTE: The “•” indicates the orientation of the filter, once the “shine” is no longer obvious.
Mix hybridization buffer with probe working solution [50ng DNA µl-1] in a 300:1 ratio.
Dip each filter completely into the Hybridization mix and place filters face-up on the parafilm covered glass slide.
Spread the rest of the solution evenly onto the filters.
Put glass slide horizontally into individual hybridization humidity chamber with corresponding % formamide to probe.
Incubate at 46°C overnight.
18:00:00
Washing: Prior Day
Make 50mL Washing Buffer in 50mL falcon tube with corresponding % formamide to probe.
Warm at 48°C overnight.
18:00:00
Washing: Day Of
Cover glass slides evenly with parafilm. Set aside until later use.
Remove filters from individual humidity chambers and put in corresponding % formamide pre-warmed Washing Buffer.
Incubate at 48°C for 10 minutes.
00:10:00
Pour 1x PBS in a Large-sized petri dish.
Transfer filters to 1x PBS and incubate for 15 minutes at room temperature.
00:15:00
Get Ice and put H2O2, Amplification Buffer, Alexa 488 in covered ice bucket.
Mix 1µL H2O2 with 199µL 1x PBS.
Create the Substrate mix in a 1000 Amplification Buffer: 10 diluted H2O2 : 3.3 Alexa 488 ratio.
Note
Eg. Substrate Mix: 1000µL Amplification Buffer, 10µL diluted H2O2, 3.3µL Alexa 488.
Note
Keep in covered ice bucket until ready to use.
Take filters out of 1x PBS and quickly wipe off excess 1x PBS on kimwipe.
Note
Make sure NOT to touch to cell-side to the kimwipe.
Dip filters into the Substrate mix and put on parafilm covered slide.
Spread the rest of the mix evenly onto the filters.
Put slides into large-sized petri dishes and seal petri dish with parafilm and put in 46°C for 45 minutes in the dark.
00:45:00
Dry filters on kimwipe and put in 1x PBS for 10 minutes at room temperature in the dark.
00:10:00
Pour MQ water into a Large-sized petri dish and pour 96% EtOH into a different Large-sized petri dish.
Transfer filters to MQ water and cover in dark for 1 min at room temperature.
00:01:00
Transfer filters to 96% EtOH and cover in dark for 1 min at room temperature.
00:01:00
Put on kimwipe on a paper towel and let dry covered in the dark.
Immediately begin resuspension step!
Note
NOTE: If for microscopy, you can store filters at -20°C.
Resuspension of Cells from Filters
Pre-heat incubator to 37°C.
Pour 1x PBS in a Large-sized petri dish.
Transfer hybridized filters to 1x PBS and incubate for 15 minutes in the dark at room temperature.
00:15:00
0.2µm filter sterilize the 150mM NaCl.
Make Resuspension Buffer (30mL 150mM NaCl; 160µL 10% Tween 80).
Put 1.5mL Resuspension Buffer per 2mL centrifuge tube and put filter in centrifuge tube.
Incubate centrifuge tube for 30 minutes at 37°C shaking horizontally.
00:30:00
Tape Tubes to vortexer horizontally (6 tubes per for vortexer).
Shake in dark for 15 minutes at 2500 rpm.
00:15:00
Remove filter (BUT keep filter just in case).
Note
The cells should now be in the buffer!!