Aug 22, 2020
5: User-friendly protocol: SABER RNA FISH in cells
This protocol is a draft, published without a DOI.
- Jocelyn Y. Kishi1,2,3,
- Sylvain W. Lapan4,3,
- Brian J Beliveau1,2,5,3,6,
- Emma R. West4,3,
- Allen Zhu1,2,
- Hiroshi M. Sasaki1,2,
- Sinem K Saka1,2,
- Yu Wang1,2,
- Constance L Cepko4,7,6,
- Peng Yin1,2,6
- 1Wyss Institute for Biologically Inspired Engineering, Harvard University, Boston, MA, USA;
- 2Department of Systems Biology, Harvard Medical School, Boston, MA, USA;
- 3These authors contributed equally;
- 4Department of Genetics, Blavatnik Institute, Harvard Medical School, Boston, MA, USA;
- 5Present address: Department of Genome Sciences, University of Washington, Seattle, WA, USA;
- 6Correspondence: py@hms.harvard.edu (P.Y.), cepko@genetics.med.harvard.edu (C.L.C.), beliveau@uw.edu (B. J. B.);
- 7Howard Hughes Medical Institute, Chevy Chase, MD, USA
- Human Cell Atlas Method Development Community
External link: http://saber.fish/
Protocol Citation: Jocelyn Y. Kishi, Sylvain W. Lapan, Brian J Beliveau, Emma R. West, Allen Zhu, Hiroshi M. Sasaki, Sinem K Saka, Yu Wang, Constance L Cepko, Peng Yin 2020. 5: User-friendly protocol: SABER RNA FISH in cells. protocols.io https://protocols.io/view/5-user-friendly-protocol-saber-rna-fish-in-cells-bh9kj94w
Manuscript citation:
Kishi, J.Y., Lapan, S.W., Beliveau, B.J. et al. SABER amplifies FISH: enhanced multiplexed imaging of RNA and DNA in cells and tissues. Nat Methods 16, 533–544 (2019). https://doi.org/10.1038/s41592-019-0404-0
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 06, 2020
Last Modified: August 22, 2020
Protocol Integer ID: 38924
Keywords: cells, SABER, RNA, Hybridization,
Abstract
This protocol describes the SABER RNA FISH in cells.
Note
Attachments
Materials
MATERIALS
Dextran sulfate, sodium saltSigma AldrichCatalog #D8906
Tween-20Sigma AldrichCatalog #P9416
µ-Slide 8 Well Glass BottomIbidiCatalog #80827
UltraPure™ DNase/RNase-Free Distilled WaterThermo FisherCatalog #10977023
UltraPure™ SSC, 20XThermo FisherCatalog #15557044
SlowFade™ Gold Antifade Mountant with DAPIThermo FisherCatalog #S36939
PBSGibco - Thermo FischerCatalog # 10010023
0.5% (vol/vol) Triton X-100
fluorescent oligos
Formamide DeionizedMerck MilliporeCatalog #S4117
Solution preparation:
Note
ddH2O is used for all solutions, not DEPC. Where possible, such as for hybridization solutions, we recommend using UltaPure DNase/RNase-free distilled water (Invitrogen #10977023).
UltraPure™ DNase/RNase-Free Distilled WaterThermo FisherCatalog #10977023
In general, we recommend good lab technique like regularly aliquoting water, 10×PBS, 20×SSC, and other solutions to avoid constantly re-opening stock bottles.
We recommend using plastic conical tubes and not lab glassware for solutions.
Formamide should be stored at 4 °C .
1×PBSTw buffer:
- 1 X PBS
- 0.1 % (v/v) Tween-20
Example mix for 1×PBSTw: 1 mL 10×PBS , 10 µL 100% Tween-20 , 9 mL H2O .
Note
Note: It is helpful to use a positive displacement pipettor to transfer Tween-20 and other detergents, but if you don’t have one available you can try to use a normal pipettor with a blunted pipette tip (cut off the bottom narrow part with scissors or a razor blade).
We recommend preparing this solution fresh the day you plan to use it.
1×PBS + Triton buffer:
- 1 X PBS
- 0.5 % (v/v) Triton X-100
Example mix for 1×PBS + Triton: 1 mL 10×PBS , 50 µL 100% Tween-20 , 9 mL H2O .
Note
We recommend preparing this solution fresh the day you plan to use it.
2×SSCT buffer:
- 2 X SSC
- 0.1 % (v/v) Tween-20
Example mix for 2×SSCT: 1 mL 20×SSC , 10 µL 100% Tween-20 , 9 mL H2O .
Note
We recommend preparing this solution fresh the day you plan to use it.
Displacement buffer:
- 1 X PBS
- 50 % to 60 % Formamide
- 0.1 % (v/v) Tween-20
Example master mix: 1 mL 10×PBS , 5 mL to 6 mL Formamide , 10 µL 100% Tween-20 , 3 mL H2O .
Note
Prepare fresh from 100 % Formamide held at 4 °C . If you are using a flow chamber, we recommend flowing displacement buffer through the chamber many times to ensure complete signal removal and also increasing to 60 % formamide as long as you are not using 30mer branches.
4×FISH master mix:
- 8 X SSC
- 40 % (wt/vol) Dextran Sulfate
- 0.4 % Tween-20
Example mix for 4×FISH master mix: 16 g Dextran Sulfate , 16 mL 20×SSC , 160 µL Tween-20 , H2O to 40 mL.
Note
Note: we recommend making 40 mL of this at a time, as it takes a while to incorporate all of the components but will last a long time. First measure 16 g Dextran Sulfate and put into a 50 mL Falcon tube. Then add the 20 X SSC and Tween-20. Add H2O to a volume of roughly 35 mL -38 g and rotate Overnight to mix all components. Finally, add H2O to adjust the final volume to 40 mL and mix again. Can be stored at Room temperature for up to several months.
Primary hybridization solution:
- 2 X SSC
- 10 % Dextran sulfate
- 0.1 % Tween-20
- 50 % Formamide
- ~100 nanomolar (nM) of each concatemer or concatemer pool
Example primary hyb mix: 31.25 µL 4×FISH master mix , 62.5 µL 100% Formamide , 8.33 µL 1µM probe 1 concatemers , 8.33 µL 1µM probe 2 concatemers , 14.59 µL ddH2O .
Note
Notes: The PER step generally produces concatemers at a concentration of 1 micromolar (µM) , and we recommend using the highest concentration you can fit into the solution to start. We often dilute 15× (to a final concentration of ~67 nanomolar (nM) ). Keep in mind if you are using a probe pool (e.g. with 50 oligos), this means the actual concentration of each strand will be much lower (1.34 nanomolar (nM) ). If you have a lot of PER concatemers you are trying to combine, or you would like to purify them, you can use a MinElute or similar column (see PER concatemerization protocol above).
We recommend creating a master mix for all of your wells (each of which should get at least 125 µL hyb ). It is helpful to use a positive displacement pipettor to transfer the 4 X FISH master mix , but if you don’t have one available you can use a normal pipettor with a blunted pipette tip (cut off the bottom narrow part with scissors or a razor blade). Mix the hyb solution very well by aggressively vortexing it for at least 00:00:10 before spinning down. Once well mixed, a normal pipettor can be used to add the hyb solution to samples, but aspiration and pipetting must be done very slowly to ensure all of the material is transferred and reduce the chance of bubble formation.
Branch hybridization solution:
- 2 X SSC
- 10 % Dextran sulfate
- 0.1 % Tween-20
- 30 % Formamide *
- 100 nanomolar (nM) ~ of each concatemer or concatemer pool
Example branch hyb mix: 31.25 µL 4×FISH master mix , 37.5 µL 100% Formamide , 8.33 µL 1µM branch 1 concatemer , 8.33 µL 1µM branch 2 concatemer , 39.59 µL ddH2O .
Note
Note: Branching is performed similarly to primary probe incubation, but in a lower formamide solution. Branches are applied after primary probe washes are complete, and before fluorescent detection. For cells, branches are extended to length of ~250-450 nt and incubated for at least 00:30:00 in 30 % formamide Hyb solution . Hyb temperature must be adjusted depending on the branches being used. We recommend using a temperature at least 1 degree lower than the lowest melting temperature of your branch sequences (see Fig. S2, protocol 3). Note that formamide concentration can be adjusted instead of oven/thermocycler temperature.
Fluorescent hybridization solution:
- 1 X PBS
- 1 micromolar (µM) each fluor oligo
Example fluorescent hyb mix: 12.5 µL 10×PBS , 12.5 µL Fluor Oligo 1 (10µM) , 12.5 µL Fluor Oligo 2 (10µM) , 87.5 µL ddH2O .
Note
Notes: We recommend starting with the 1 hour hybridization with 1 micromolar (µM) fluor oligo if this is your first time running the protocol. Alternatively, we still see strong signal if the hybridization time is reduced to 00:15:00 (at Room temperature e.g. for Fig. 6 (see Nature Methods article)). Side-by-side testing with two probes in tissue indicate 0.2 micromolar (µM) fluorescent oligo is also sufficient (see Fig. S3, protocol 6).
Safety warnings
For hazard information and safety warnings, please refer to the SDS (Safety Data Sheet).
Before start
Preparing the slide
Preparing the slide
Seed 8-well Ibidi chamber slides (Cat #80827) with cells and grow in tissue culture incubator (typically 37 °C with 5 % CO2 ) to desired confluency.
Note
Notes: You may need to adjust the deposition protocol (e.g. what concentration of cells, how long to let them grow on the slide) in order to achieve the desired confluency of your cell type for imaging.
Some cell types will not adhere well through the fixation protocol, so the chamber should be quickly imaged in Brightfield after fixation to ensure the proper density of cells.
If this is your first time doing this protocol, we recommend using some of the wells for controls and a couple probe length conditions. For example, you might seed 4 wells and use one as a no probe control well (which receives all other treatments) where you should not see signal, and then 3 different concatemer length conditions. Or if you have 3 targets, you might do 3 single-color wells each receiving one of the concatemer species, and then one multi-color well. We have found that some microscope stages are not amenable to imaging the leftmost and rightmost wells in the chamber, so this is a good thing to check before you plan on using all of the wells.
Depending on the type of fluid, the minimum volume for each well is around 120 µL -150 µL . We recommend using 250 µL for washes. If you are using a different type of chamber with smaller volume, we recommend performing several more washes at each step to ensure complete removal of previous elements. The easiest method of aspiration is with an unfiltered pipette tip attached to a vacuum line. We recommend cleaning the line with ethanol and changing the tip a several times throughout the experiment. Good lab technique for avoiding RNase contamination and use of RNase-free water is important throughout the protocol.
Fixation
Fixation
At Room temperature , rinse cells in 1 X PBS and then immediately fix in 4 % (wt/vol) paraformaldehyde for 00:10:00 .
Rinse again in 1 X PBS and store at 4 °C .
Note
We stored chambers for DNA FISH up to a couple of weeks before use, but for RNA integrity we recommend waiting no longer than a few days.
Hybridization
Hybridization
Wash in 1 X PBS for 00:01:00 at Room temperature .
Permeabilize in 1 X PBS + 0.5 % Triton X-100 for 00:10:00 at Room temperature .
Wash in 1 X PBSTw for 00:01:00 at Room temperature .
Wash in 2 X SSCT for 00:01:00 at Room temperature .
Add hybridization solution and denature at 60 °C for 00:03:00 .
Return temperature to 42 °C for at least 04:00:00 (typically Overnight is easiest).
Note
Note: if the probes are retrieved using the Oligominer33 pipeline as described or from the Wu lab database, then 42 °C hyb temperature should be sufficient for RNA FISH. If you are using custom designed and especially shorter probes, you will need to check their melting temperatures in 2 X SSC + 50 % formamide . We suggest setting your oven temperature to a couple degrees lower than those melting temperatures (See protocol 3). We have found that thermocyclers especially with heated lids versus ovens set to the same temperature can actually be up to a few degrees higher at the sample plane. This can cause a big difference in yield because we are operating close to the melting temperatures, so if you feel that your signal is too low you can try reducing the temperature of the hyb a bit. Alternatively, if there seems to be a lot of off target binding of probes, you can try increasing a bit.
Add 200 µL pre-warmed 2 X SSCT to wells and aspirate.
Note
Note: For pre-warming 2 X SSCT , we suggest you first make enough for all washes and then aliquot the amount needed for the hot washes into 2 mL tubes. Put these tubes on a heated tube rack set to 65 °C and allow them to heat for at least 00:30:00 -00:40:00 . Remove only the amount of tubes you need at each wash step, so that the rest can stay hot.
Wash 4×00:05:00 in pre-warmed 2 X SSCT at 60 °C .
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (1/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (2/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (3/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (4/4)
Wash 2×00:02:00 in 2 X SSCT at Room temperature .
Wash 00:02:00 in 2 X SSCT at Room temperature . (1/2)
Wash 00:02:00 in 2 X SSCT at Room temperature . (2/2)
If you are going directly to the fluorescent protocol, rinse in 1 X PBS for 00:01:00 and transfer to fresh 1 X PBS (atRoom temperature ).
Pause point: sample can be stored at 4 °C Overnight to several days.
Branching
Branching
Wash in 2 X SSCT for 00:02:00 at Room temperature .
Add branch hybridization solution and hold at 37 °C for at least 00:30:00 .
Note
Note: Branching is performed similarly to primary probe incubation, but in a lower formamide solution. Branches are applied after primary probe washes are complete, and before fluorescent detection. For cells, branches are extended to length of ~250-450 nt and incubated for at least 30 minutes in 30 % formamide Hyb solution . Hyb temperature must be adjusted depending on the branches being used. We recommend using a temperature at least 1 degree lower than the lowest melting temperature of your branch sequences (see Fig. S2, protocol 3). Note that formamide concentration can be adjusted instead of oven/thermocycler temperature.
Add 200 µL pre-warmed 2 X SSCT to wells and aspirate.
Wash 4×00:05:00 in pre-warmed 2 X SSCT at 60 °C .
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (1/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (2/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (3/4)
Wash 00:05:00 in pre-warmed 2 X SSCT at 60 °C . (4/4)
Wash 2×00:02:00 in 2 X SSCT at Room temperature .
Wash 00:02:00 in 2 X SSCT at Room temperature . (1/2)
Wash 00:02:00 in 2 X SSCT at Room temperature . (2/2)
Rinse in 1 X PBS for 00:01:00 .
Transfer to fresh 1 X PBS (at Room temperature ).
Fluorescent detection
Fluorescent detection
Rinse once in 1 X PBS at Room temperature .
Add fluor hybridization solution and hold at 37 °C for 01:00:00 .
Note
Notes: We recommend starting with the 1 hour hybridization with 1 micromolar (µM) fluor oligo if this is your first time running the protocol. Alternatively, we still see strong signal if the hybridization time is reduced to 00:15:00 (at Room temperature e.g. for Fig. 6 in the Nature Methods article). Side-by-side testing with two probes in tissue indicate0.2 micromolar (µM) fluorescent oligo is also sufficient (see Fig. S3, protocol 6).
Wash for 00:05:00 with pre-warmed 1 X PBS at 37 °C .
Note
Note: For pre-warming 1 X PBS , we suggest you first make enough for all washes and then aliquot the amount needed for the warming washes into a separate Falcon tube and place into an oven or warm room at 37 °C - 45 °C .
Wash 2×00:02:00 in pre-warmed 1 X PBS .
Wash 00:02:00 in pre-warmed 1 X PBS . (1/2)
Wash 00:02:00 in pre-warmed 1 X PBS . (2/2)
Rinse once in 1 X PBS at Room temperature .
Load SlowFade + DAPI (Thermo Fisher S36939) mountant to completely cover cells.
Note
Note: You can also image samples directly in 1 X PBS , in which case if you may need to do a short DAPI stain if you want to see that channel.
Image.
Pause point: Samples can be stored at 4 °C before imaging for a few days, although we recommend doing the fluorescent hyb closer to your imaging session time (day of) where possible.
Serial detection (complete before repeating fluorescent detection)
Serial detection (complete before repeating fluorescent detection)
Wash 2×00:01:00 in PBS at Room temperature to remove mountant.
Wash 00:01:00 in PBS at Room temperature to remove mountant. (1/2)
Wash 00:01:00 in PBS at Room temperature to remove mountant. (2/2)
Add displacement buffer and incubate for 00:15:00 at Room temperature .
Note
Notes: the signal should be stripped almost immediately with the displacement buffer, but we recommend letting it sit for at least 00:05:00 -00:15:00 to ensure complete removal. If you are using a smaller volume chamber or well, you may need to add fresh displacement buffer extra times in order to completely strip fluorescent signal. If you are doing the exchange in place on a microscope, you can image the same area before and after adding the displacement buffer to visualize the drop in signal. If you are using DAPI, the displacement buffer will probably remove most of this signal. You will likely need to replace after each exchange step and before imaging, either by including it in the mountant or staining separately (see above).