TITLE: Fluorescent In Situ Hybridization (FISH - RNAscope) in mouse brain sections
AUTHORS: Yu Lin Tan, Oriol Pavon Arocas, Lucille Duquenoy, Tiago Branco

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block">Here we describe a protocol to perform multiplex fluorescent in situ hybridization (FISH) in thin sections from frozen murine brain tissue. The protocol can be divided in several days or can be completed in two successive days: one day for extracting and slicing the brain and a second one for the FISH.</div></div>

[STEPS]
?. [Day 0 | Brain extraction]
[Preparation ~ 15 min]
You can extract and freeze several brains on the same day and then store them at -80°C until sectioning. You can do the brain extraction either in the PFA perfusion room (with tools from there) or in the slice electrophysiology area where you prepare acute brain slices (with clean tools). Never bring tools contaminated with PFA to the electrophysiology area.
?. [Day 0 | Brain extraction]
You will need:A small bucket or ice pan with dry ice.A bigger tray with wet ice.A plastic bag for the carcass.Tools for decapitation and brain extraction: big scissors for decapitation, medium scissors to cut through skin, small scissors to cut skull, forceps to separate skull, spatula to remove the brain, brush to help position the brain.One mould for each brain (this is where you will freeze the brains in OCT to make a little cube you will later slice on the cryostat).2-methylbutane (from the chemicals cabinet in the Histology area of Level 1) - It speeds up freezing in the dry ice.A ~250ml glass bottle with 1x PBS per brain.Filter paper.1x phosphate-buffered saline (PBS, pH 7.4) .Glassware: a ~150mL beaker where you will put PBS (to clean the blood from the brain), a glass petri dish where you will put PBS (to clean the blood from the brain).
Further details for the tools and steps related to brain extraction can be found on Oriol Pavon Arocas, Tiago Branco (2021). Preparation of acute midbrain slices for patch-clamp recordings. [https://dx.doi.org/10.17504/protocols.io.bukhnut6]
?. [Day 0 | Brain extraction]
[Decapitation under terminal anaesthesia and Brain extraction ~ 15 min per brain]
You can do this following two paths: after terminal anaesthesia via intraperitoneal (IP) injection of euthatal, or after terminal isoflurane anaesthesia (as we do for acute brain slices).
?. [Day 0 | Brain extraction]
[Common path]Label each mould with the animal ID, name of experimenter, and date beforehand. Label in a consistent manner, so you know where the top, down, right, and left of the brain are.Fill the moulds with OCT, very carefully to avoid bubbles. Keep them flat on ice on the tray.Put a bit of dry ice in the ice pan. Don't put too much as you will need to make a pocket for the mould to stand flat in the floor of the ice pan.Fill beakers with cold PBS and also leave on ice on the tray.
?. [Day 0 | Brain extraction]
[Bifurcation - PATH A] - in Fume Hood of Perfusion RoomOverdose by IP injection of euthatal - Decapitation - Brain extractionFirst, get and prepare the drugs you will need from the surgery prep room: we use euthatal diluted in saline. Always follow the most recent animal welfare guidelines of your home institution. Mix in a bijou vial and load on an insulin needle syringe.Wear appropriate PPE for the perfusion room (hair net, lab coat, gloves, mask, have fume hood on).Once everything is ready, proceed to scruff and inject the mouse. Wait until the anaesthesia has its effect and check reflexes to confirm the animal is fully anaesthetised.Move to the hood: put the tray in the hood. Double check your tools are ready (specific for perfusion room, don't mix with the clean ones for slice electrophysiology).Decapitate under the hood and extract the brain as you would do for acute brain slices (see https://dx.doi.org/10.17504/protocols.io.bhfzj3p6 for a step by step description). Briefly, drop the head in a beaker with PBS to clean the blood, cut through the skull, extract the brain, and carefully drop the brain in the small petri dish with PBS to wash the blood away.
?. [Day 0 | Brain extraction]
[Bifurcation - PATH B] - in Slice Electrophysiology AreaTerminal anaesthesia with isoflurane - Decapitation - Brain extractionMove to bench to extract the brain as you would do for acute brain slices (see https://dx.doi.org/10.17504/protocols.io.bhfzj3p6 for a step by step description). Carefully drop the brain in the small petri dish with PBS to wash the blood away.
?. [Day 0 | Brain extraction]
[Back to common path]Carefully lift the brain with the spatula from the PBS and transfer it to a piece of filter paper.With the help of a brush and the spatula, dry as much PBS from the brain as possible - this is important to ensure that the OCT sticks to the brain directly. [!] Careful - the brain will be soft and will tend to stick to the filter paper when dry, do not over dry it as you will risk damaging the tissue.Once dry, transfer the brain to the mould and immerse it in OCT.Be consistent when placing the brain in the mould: we always put the olfactory bulb down, with the dorsal part of the brain facing upwards (with respect to the written edge). In the moulds we use, there are two tiny ridges on one of the walls - we make that the dorsal wall, the one where cortex points to. Use the small forceps to very carefully arrange the brain as flat and as centered as possible (it is very important to displace as little OCT as possible - and avoid creating bubbles!).Finally, add a bit of 2-methylbutane on the dry ice (this will speed up freezing) and make a bit of space for the mould. Transfer the mould to the dry ice, making sure the mould sits flat on the bottom of the ice pan and is surrounded by dry ice.You can now proceed to extract the next brain - by the time you are ready to freeze it, the previous mould will already be frozen and you will be able to move it to a corner of the ice pan and leave it on dry ice until you complete the procedure for all the brains.Once you are done and all the brains are frozen in their moulds with OCT, you can put them in a labeled plastic zip bag and store them at the -80°C freezer until cryosectioning.Use the small forceps to very carefully arrange the brain as flat and as centered as possible (it is very important to displace as little OCT as possible - and avoid creating bubbles!).Finally, add a bit of 2-methylbutane on the dry ice (this will speed up freezing) and make a bit of space for the mould. Transfer the mould to the dry ice, making sure the mould sits flat on the bottom of the ice pan and is surrounded by dry ice.You can now proceed to extract the next brain - by the time you are ready to freeze it, the previous mould will already be frozen and you will be able to move it to a corner of the ice pan and leave it on dry ice until you complete the procedure for all the brains.Once you are done and all the brains are frozen in their moulds with OCT, you can put them in a labeled plastic zip bag and store them at the -80°C freezer until cryosectioning.
?. [Day 0 | Brain extraction]
[!] PAUSE point - once the brain is frozen and kept at -80°C you can slice it the next day, week, or month.
?. [Day 1 | Cryostat Sectioning]
[!] Make sure you book the cryostat in advance if shared with others. Some of the following steps will vary depending on the cryostat model in your institute. The total time will depend on the number of brains you are doing.
?. [Day 1 | Cryostat Sectioning]
[Preparation - tools]First, collect your brains from the -80°C freezer and leave the moulds at the cryostat to equilibrate to -20°C for one hour.In the meantime, get what you will need:SuperFrost slides - Usually have 3 series (labelled A1, B1, C1) so that you add one section to each to end up with every third section in any given slide.Write key information with pencil (Animal, date, etc.).Cryostat blade (reusable, usually kept in a falcon tube).OCT from histology area.A slide box (for storage at -80°C).2 small brushes and 1 big brush (from the cryostat top).
?. [Day 1 | Cryostat Sectioning]
[Preparation - cryostat]Set the cryostat temperature at -16°C for CT and -17°C for OT - this may very depending on your model and room temperature.You may have to play with the temperature during cutting: if sections break, then the temperature is too low and you can try to bring it up by opening the sliding cover a bit more.When you are done leave at -14°C to save energy.If screen/controls don't work or say Locked, press the Key/Lock button for several seconds until it unlocks.
?. [Day 1 | Cryostat Sectioning]
[Mount brain for sectioning]Place the blade on the holder.Given that we reuse them, try to offset it a bit so you don't always blunt the same region.Put some tissue paper beneath the blade to make it easier to clean up.Get the round holder and add some OCT in the center.Break the molds, take the frozen brain out of the mold and press the cube of frozen OCT with the brain against the blob in the round holder: the OCT will spread out and act as glue.Put it inside the cryostat and leave to freeze.Once frozen, mount the holder with the brain on the arm of the cryostat.Unlock it and adjust so that it is flat ("zero it").
?. [Day 1 | Cryostat Sectioning]
[Trim until you reach the area of interest]Set the cutting thickness to 30 μm (or your desired thickness) and the trimming one to 100 μm.On trimming mode, remove the "lock" from the wheel on the side and start cutting and keep adjusting so the sections come out flat and even.
?. [Day 1 | Cryostat Sectioning]
[Collect brain sections]When you reach your area of interest, use a razor blade to cut out as much OCT as possible so that when you mount them on the slide the OCT doesn't take extra space. It is a trade-off and you need to be careful - if you remove too much of it the sections will not hold.[IMP!] For FISH it is critical that the tissue touches the glass directly, if it ends up on top of a bit of OCT, you will loose the sections during the washes of the protocol.Slice carefully, don't go fast.Collect the sections making sure they don't wrinkle/fold on themselves.Once on the slide, keep outside so the OCT melts and then dries out.When you are done collecting sections or fill up one slide, leave it out so the OCT dries out, then put in the box and keep in the chamber at -20°C until the end.
?. [Day 1 | Cryostat Sectioning]
[!] PAUSE point - Once sectioning is completed and all the slides are dry and in the storage box, store the box at -80°C until you are ready to do the FISH protocol.
?. [Day 2 | FISH: fixation and dehydration]
In preparation for this step, defrost 4% PFA (you need ~50 ml to fill one glass slide holder) to 4 degrees or prepare fresh 4% PFA from 16% PFA vials (dilute 12.5 ml and bring to 50 ml with PBS).In addition, think of which and how many slides you will need for each experiment, and always take an extra one for the positive and negative controls.
This step includes work with paraformaldehyde (PFA), make sure to follow all safety protocols in place at your institution.
?. [Day 2 | FISH: fixation and dehydration]
[Fixation ~ 1 hour]Take the selected slides out of the -80°C freezer and immediately place into PFA (put the slides into the glass slide holder and then pour PFA along the sides).[IMP!] always pour the solution (be it PFA, PBS or wash buffer) along the sides that the sections are not facing, or you may accidentally remove them from the slide.Fix at 4 degrees for 1 hour.Remove the PFA (dispose appropriately) and perform 2x quick washes in PBS by adding PBS, moving slides up and down 3-5 times (washes should be quick), and repeating after replacing PBS with fresh one.
?. [Day 2 | FISH: fixation and dehydration]
[Dehydration ~ 20 min]In three different falcon tubes, freshly prepare 50 ml of 50% ethanol, 50 ml of 70% ethanol, and 50 ml of 100% ethanol by diluting in pure ddH20.Place slides in 50% ethanol for 5 min at Room Temperature (RT) - i.e. pour out the PBS from the glass slide holder and pour in the 50ml of ethanol.Repeat with 70% ethanol, 5 min, RT.Repeat with 100% ethanol, 5 min, RT.
?. [Day 2 | FISH: fixation and dehydration]
[OPTIONAL - Store at -20°C for up to 1 day]Remove the ethanol from the holder and add 50 ml of fresh 100% ethanol to cover the slides.Wrap the glass slide holder with cling film (or parafilm) to prevent ethanol from evaporating and store at -20°C for up to 1 day (it is best to start subsequent steps by the next day).
This step is optional - you can ignore this step and proceed to subsequent steps directly.
?. [Day 3 | FISH: staining and mounting]
Before starting this part, make sure you have planned the combination of probes and channels you will use (which will determine the colors to image - see note at the end).
?. [Day 3 | FISH: staining and mounting]
[Bake slides at 40 degrees ~ 15-20 min]Preheat the RNAscope oven to 40°C.Remove slides from slide glass holder and let ethanol evaporate. Wipe the bottom side with a clean wipe.While drying, cut parafilm to exactly the size of each of your sample areas. The parafilm will prevent evaporation of the reagents and ensure they evenly cover all the brain sections.Place slides directly on tray (without rack).Bake slides at 40 degrees for 15-20 min (the longer you bake them the better the sections will stick to the slide).In the meantime, warm water bath to 40°C and bring Protease IV vial to RT.[!] Make sure you take Protease IV (and not III).
?. [Day 3 | FISH: staining and mounting]
[Draw hydrophobic barrier ~ 10 min, depending on number of slides]Use the Immedge pen - lines should be blue when drawn (do not use the PPAP pen, which draws green lines).[!] Draw on multiple layers - it is very important to ensure reagents stay on the slide.Draw on three of the sides (leave one blank to remove excess liquid).
?. [Day 3 | FISH: staining and mounting]
[Apply Protease IV at RT and prepare probe mixtures ~ 30 min]Place the slides on the black slide tray from Sigma (this is only so that you can see the sections - you can use the baking tray or tissue paper if you don't have it).Add ~4-5 drops/slide of Protease IV (ensure there is enough and always err on the excess side - you don't want the slides to dry out).Place parafilm on, ensuring there are no bubbles (remember you previously cut the parafilm to exactly the size of your sample area). [! VERY IMPORTANT STEP !]While waiting during the 30 min, prepare the reagents for the first hybridization step:Dilute 20 ml of 50X Wash Buffer by adding ddH2O up to 1 L.Warm up probes (entire bottle) for 10 min at 40°C in the water bath, then let cool to RT.Briefly spin probe bottles.Prepare probe mixture in 2 ml eppies - 50:1:1 ratio of C1:C2:C3 probes (you will need to plan the combination in advance - see note at the end).Use 150μl : 3μl : 3μl per slide until you are very confident with the protocol, then reduce to 100:2:2 per slide.Warm the HybEZ slide rack in the oven to 40°C.After 30 min, lift the parafilm and shake off excess reagent (there should be excess reagent to shake off!! Else insufficient reagent was applied or barrier was not done well).Dab the parafilm dry - make sure you keep the same piece for the same slide, especially if you use different probe/control combinations.Perform 2x quick washes in PBS by moving slides up and down 3-5 times (washes should be quick).Dab excess liquid off (blot onto tissue from side and back of slide).Place some wet tissue paper on the bottom of the HybEZ slide rack to keep it humid for the next steps.While waiting during the 30 min, prepare the reagents for the first hybridization step:Dilute 20 ml of 50X Wash Buffer by adding ddH2O up to 1 L.Warm up probes (entire bottle) for 10 min at 40°C in the water bath, then let cool to RT.Briefly spin probe bottles.Prepare probe mixture in 2 ml eppies - 50:1:1 ratio of C1:C2:C3 probes (you will need to plan the combination in advance - see note at the end).Use 150μl : 3μl : 3μl per slide until you are very confident with the protocol, then reduce to 100:2:2 per slide.Warm the HybEZ slide rack in the oven to 40°C.After 30 min, lift the parafilm and shake off excess reagent (there should be excess reagent to shake off!! Else insufficient reagent was applied or barrier was not done well).Dab the parafilm dry - make sure you keep the same piece for the same slide, especially if you use different probe/control combinations.Perform 2x quick washes in PBS by moving slides up and down 3-5 times (washes should be quick).Dab excess liquid off (blot onto tissue from side and back of slide).Place some wet tissue paper on the bottom of the HybEZ slide rack to keep it humid for the next steps.After 30 min, lift the parafilm and shake off excess reagent (there should be excess reagent to shake off!! Else insufficient reagent was applied or barrier was not done well).Dab the parafilm dry - make sure you keep the same piece for the same slide, especially if you use different probe/control combinations.Perform 2x quick washes in PBS by moving slides up and down 3-5 times (washes should be quick).Dab excess liquid off (blot onto tissue from side and back of slide).Place some wet tissue paper on the bottom of the HybEZ slide rack to keep it humid for the next steps.
?. [Day 3 | FISH: staining and mounting]
[Hybridize probes at 40 degrees ~ 2 hours]Hybridize probes at 40°C for 2 hours - Add ~150 μl/slide and place parafilm without bubbles.First place the slides on the HybEZ slide rack. Then add the probe mixtures to the corresponding slides and cover with parafilm without bubbles. Only then secure the slides on the rack. Finally, put the tray in the oven and leave to incubate for 2h. Repeat this procedure on the following steps.Once the time is up, lift parafilm, dab excess liquid off, and shake off excess reagent from the slide (there should be excess reagent to shake off!! Else insufficient reagent was applied, barrier was not done well, or chamber was not humid enough).Perform 2x 2 min washes in Wash Buffer at RT (normal washes from now on, not the quick ones). After the second wash is done, shake off excess liquid (blot onto tissue from side and back of slide) and dry the bottom of the slides before proceeding to the next step.
?. [Day 3 | FISH: staining and mounting]
[Hybridize Amp1-FL at 40 degrees ~ 30 min]Hybridize Amp1-FL at 40°C for 30 min - Add ~4-5 drops/slide and place parafilm without bubbles.Once the time is up, lift parafilm, dab excess liquid off, and shake off excess reagent from the slide (there should be excess reagent to shake off!! Else insufficient reagent was applied, barrier was not done well, or chamber was not humid enough).Perform 2x 2 min washes in Wash Buffer at RT (normal washes, not the quick ones). After the second wash is done, shake off excess liquid (blot onto tissue from side and back of slide) and dry the bottom of the slides before proceeding to the next step.
?. [Day 3 | FISH: staining and mounting]
[Hybridize Amp2-FL at 40 degrees ~ 15 min]Hybridize Amp2-FL at 40°C for 15 min - Add ~4-5 drops/slide and place parafilm without bubbles.Once the time is up, lift parafilm, dab excess liquid off, and shake off excess reagent from the slide (there should be excess reagent to shake off!! Else insufficient reagent was applied, barrier was not done well, or chamber was not humid enough).Perform 2x 2 min washes in Wash Buffer at RT (normal washes, not the quick ones). After the second wash is done, shake off excess liquid (blot onto tissue from side and back of slide) and dry the bottom of the slides before proceeding to the next step.
?. [Day 3 | FISH: staining and mounting]
[Hybridize Amp3-FL at 40 degrees ~ 30 min]Hybridize Amp3-FL at 40°C for 30 min - Add ~4-5 drops/slide and place parafilm without bubbles.Once the time is up, lift parafilm, dab excess liquid off, and shake off excess reagent from the slide (there should be excess reagent to shake off!! Else insufficient reagent was applied, barrier was not done well, or chamber was not humid enough).Perform 2x 2 min washes in Wash Buffer at RT (normal washes, not the quick ones). After the second wash is done, shake off excess liquid (blot onto tissue from side and back of slide) and dry the bottom of the slides before proceeding to the next step.
?. [Day 3 | FISH: staining and mounting]
[Hybridize Amp4-FL at 40 degrees ~ 15 min]Hybridize Amp4-FL at 40°C for 15 min - Add ~4-5 drops/slide of the correct [!] Amp4 (either A, B, or C) depending on your chosen color combination.Once the time is up, lift parafilm, dab excess liquid off, and shake off excess reagent from the slide (there should be excess reagent to shake off!! Else insufficient reagent was applied, barrier was not done well, or chamber was not humid enough).Perform 2x 2 min washes in Wash Buffer at RT (normal washes, not the quick ones). After the second wash is done, shake off excess liquid (blot onto tissue from side and back of slide) and dry the bottom of the slides before proceeding to the next step.
?. [Day 3 | FISH: staining and mounting]
[DAPI at RT ~ 3 min]Apply DAPI at RT for 3 min.Or, alternatively, use mounting media with DAPI and skip this step.
?. [Day 3 | FISH: staining and mounting]
[Mount slides ~ 1 hour 15 min, depending on number of slides]Dry excess DAPI off (without washing!).Mount slides:[!] Clean the coverslips with ethanol and let dry.[!] Use a tissue to carefully dry as much liquid around the sections as you can. Then let them dry completely before mounting (around 1 hour). Otherwise possible artefacts can appear when imaging.Once dry, add mounting media (with or without DAPI). Do not add in excess - avoid there is excess media coming out the edges of the coverslip.Carefully place the coverslip without creating bubbles and apply nail polish to seal.Keep in a box at 4°C until imaging.Mount slides:[!] Clean the coverslips with ethanol and let dry.[!] Use a tissue to carefully dry as much liquid around the sections as you can. Then let them dry completely before mounting (around 1 hour). Otherwise possible artefacts can appear when imaging.Once dry, add mounting media (with or without DAPI). Do not add in excess - avoid there is excess media coming out the edges of the coverslip.Carefully place the coverslip without creating bubbles and apply nail polish to seal.Keep in a box at 4°C until imaging.
?. [Choosing your probe combination]
[Choosing your channels appropriately]*Source: https://acdbio.com/rnascope%C2%AE-fluorescent-multiplex-assay.
[IMP] Make sure you plan the combination of probes and channels you will use in advance (which will determine the colors to image and the Amp4 you will need to use).
?. [Choosing your probe combination]
Channel selection for multiplexing:Guidelines for Fluorophore Assignment:Fluorescein - Green
For Alexa Fluor 488Visible with naked eyeLeast distinct from most tissue autofluorescenceHigh ExpressorCyanine 3 - Orange
For Atto 550Visible with naked eyeNoneLow Expressor or Expression Level UnknownCyanine 5 - Far Red
For Atto 647NEasily differentiated from autofluorescenceNot visible to the naked eyeLow Expressor
Fluorescein - Green
For Alexa Fluor 488Visible with naked eyeLeast distinct from most tissue autofluorescenceHigh ExpressorCyanine 3 - Orange
For Atto 550Visible with naked eyeNoneLow Expressor or Expression Level UnknownCyanine 5 - Far Red
For Atto 647NEasily differentiated from autofluorescenceNot visible to the naked eyeLow Expressor
?. [Choosing your probe combination]
[Positive and Negative Controls]*Source: https://acdbio.com/control-slides-and-control-probes-rnascope.Control Probes for Manual Assays - RNAscope Multiplex Fluorescent Reagent Kit:Species-specific positive probes and negative control probes are needed to run with human or mouse control slides for quality control check of the technique and with your sample for the Sample or RNA quality control check. ACD provides positive control probes that target species-specific housekeeping genes and negative control probes that target the bacterial DapB gene.Careful selection of the right positive control is recommended. The expected expression level of your target RNA is the key criteria to consider as your positive control probe should have a similar expression profile.When working with RNAscope Multiplex Fluorescent Reagent Kit, you will have ultimate flexibility and can use with one, two or three target RNAs.When using this RNAscope Multiplex Fluorescent Reagent Kit with three target RNAs, you will work in channels 1, 2 and 3 so you will also need control probes for channel 1, channel 2 and channel 3. These species-specific positive control probes and negative control probes for channel 1 when processed with your sample will enable you to assess your sample/RNA quality and serve as control for your channel 1 target probe etc. ACD offers a 3-plex Negative Control Probe which contains probes for channel 1, 2 and 3. We also offer 3-plex RNAscope® Positive Control Probes for Human, Mouse and Rat where POLR2A probe is available in channel 1 and PPIB probe in channel 2 and UBC probe in channel 3.
?. [Choosing your probe combination]
Positive control:Control probes are already mixed in a bottle labelled "3-plex Positive Control Probe", Cat. No. 320881, website link: https://acdbio.com/search/site/%252A320881%252A/cms/probes.RNAscope Mouse positive control probe for RNAscope Multiplex Fluorescent Assay: Polr2a (C1 channel) and PPIB (C2 channel), UBC (C3 channel).Note: In terms of relative expression levels, UBC is the highest, PPIBis considered a moderate-high, and POLR2A is a moderate-to-low expressor target.The positive control will allow you to identify whether the lack of signal for a probe is due to the target not being expressed or due to a problem in the reagent kit you used: if the empty channel is both in your "test" sections and your "positive control" sections, then it is likely that one (or more) of the reagents or steps did not work.
?. [Choosing your probe combination]
Negative control:Control probes are already mixed in a bottle labelled "3-plex Negative Control Probe", Cat. No. 320871, website link: https://acdbio.com/search/site/%252A320871%252A/cms/probes.