The MERSCOPE is an automated platform for the acquisition of high-quality MERFISH data for spatially resolved single-cell transcriptomics. In this video, we will walk you through a general protocol for preparing fresh and fixed frozen tissue samples for analysis . For the complete protocol, please refer to the applicable User Guides located in the Resources section of our website"

Begin by preparing a cryosection of either fresh or fixed frozen tissue of interest. Tissue blocks should be embedded with optimal cutting temperature, or OCT, compound and stored at minus 80 to prevent tissue drying. Place the tissue block into the cryotome and allow it to sit at minus 20 degrees C for at least 30 minutes before sectioning. Always use new blades when sectioning tissue. Trim the block for the desired tissue region and cut a 10-micrometer section. When trimming to expose the tissue of interest, discard at least the first 50 μm slices of the tissue prior to sectioning for MERSCOPE samples .
Carefully lower the MERSCOPE Slide onto the tissue section, ensuring the tissue section is flat and mounted in the centre of the slide. Let it sit for about 5 seconds. The OCT compound will turn white as the tissue refreezes. Now place the slide face up onto a dry, labeled petri dish and let it sit in minus 20 degrees Celsius for 5 minutes. This will allow the tissue to adhere to the slide .
Begin by cleaning the lab bench and tools with RNase cleaning solution. For fixation of fresh frozen tissue, perform in a fume hood. Add 5 milliliters of fixation buffer to the slide. And incubate at room temperature for 15 minutes. This step is not necessary for fixed frozen tissue.

For tissue permeabilization, begin by washing the slide with 5 milliliters of 1X PBS. Incubate for 5 minutes and aspirate the buffer. Repeat this a total of 3 times. Finally, add 5 milliliters of 70% ethanol. Seal the petri dish with parafilm and place at 4 degrees Celsius overnight to let the tissue permeabilize.

If necessary, the sample can be placed in the MERSCOPE Photobleacher for autofluorescence quenching. Cell Boundary Staining may also be performed to better mark the boundaries between individual cells.

Place the parafilm-sealed petri dish under the MERSCOPE Photobleacher. ENSURE there are no labels, writing, other items on the lid that may block the light. Turn on the MERSCOPE Photobleacher and leave at room temperature for at least 3 h. Autofluorescence quenching may also be performed when the sample is stored in Clearing Solution .

II. Cell Boundary Staining
Prepare blocking solution. Aspirate the 70% ethanol and wash with 5 mL 1X PBS.
Aspirate the 1X PBS to dry the MERSCOPE Slide, leaving just enough liquid to cover the tissue section.
Add 100 μL of Blocking Solution onto the center of the tissue section. Use scissors to cut a piece of parafilm 2×2 cm. Use tweezers to peel off the parafilm backing and place the side previously protected by the backing onto the solution. Avoid introducing air bubbles. Incubate at room temperature for 1 h.

Prepare Primary Staining Solution. Use tweezers to remove the parafilm square from the tissue section. Aspirate the solution to dry the MERSCOPE Slide, leaving just enough liquid to cover the tissue section.

Add 100 μL Primary Staining Solution onto the center of the tissue section. Use scissors to cut a piece of parafilm 2×2 cm. Use tweezers to peel off the parafilm backing and place the side previously protected by the backing onto the solution. Avoid introducing air bubbles. Incubate at room temperature for 1 h.

Use tweezers to remove the parafilm from the tissues section and wash with 5 mL 1X PBS. Incubate for 5 min on a rocker. Repeat this for a total of 3 times.

Prepare Secondary Staining Solution.
Aspirate the 1X PBS to dry the MERSCOPE Slide, leaving just enough liquid to cover the tissue section. Add 100 μL Secondary Staining Solution onto the center of the tissue section. Use scissors to cut a piece of parafilm 2×2 cm. Use tweezers to peel off the parafilm backing and place the side previously protected by the backing onto the solution. Avoid introducing air bubbles. Incubate at room temperature for 1 h.

Use tweezers to remove the parafilm square from the tissue section and wash with 5 mL 1X PBS. Incubate for 5 min on a rocker. Repeat this for a total of 3 times.

Aspirate the 1X PBS. In a fume hood, add 5 mL fixation buffer to fix the stained tissue section at room temperature for 15 min. Wash with 5 mL 1X PBS and incubate for 5 min at room temperature. Repeat this two times. Proceed immediately to the next step

Begin the Encoding Probe Hybridization step by aspirating any remaining solution from the dish containing the MERSCOPE slide and tissue section.

Wash with 5 milliliters of sample prep wash buffer, then aspirate.

Add 5 milliliters of formamide wash buffer and cover the dish. Be sure to use the fume hood when handling formamide.

Incubate at 37 degrees Celsius for 30 minutes.

After incubation, aspirate the formamide wash buffer to dry the region of the slide outside of the tissue section .

Add 50 milliliters of MERSCOPE Encoding Probe set directly onto the center of the tissue section.

Cut out a 2-by-2 centimeter strip of parafilm. Using tweezers, peel off the backing and gently place the protected side of the parafilm onto the solution.

Lift and lower the parafilm so that the encoding probe set is spread across the entire tissue section. Avoid any air bubbles and make sure that the parafilm fits within the slide, otherwise the probe set may wick out into the petri dish.

Place the petri dish into a larger dish and cover. Spray the outside with 70% ethanol to sterilize.

Then incubate in a humidified 37 degree C cell culture incubator for at least 36 hours, or a maximum of 48 hours.

After incubation, remove the parafilm. Add 5 milliliters of Formamide wash buffer.

Using a mini-incubator in a fume hood, incubate the sample at 47 degrees C for 30 minutes.

Aspirate the formamide wash buffer.

Wash 2 times with 5 milliliters of Sample Prep Wash Buffer.

We will now proceed with Gel Embedding. Using tweezers, take out one Gel Coverslip.

Clean it by spraying with RNaseZap solution and wiping with a Kimwipe.

Then spray with 70% ethanol and wipe again.

Add 50 microliters of Gel Slick Solution onto the gel coverslip.

Spread the Gel Slick by gently wiping with a Kimwipe.

Prepare the Gel Embedding Solution.

Aspirate the Sample Prep Wash buffer.

Add 5 milliliters of Gel Embedding Solution. Incubate at room temperature for 1 minute.

Transfer a majority of the Gel Embedding Solution to a waste tube to monitor the gel formation.

Aspirate out the remaining solution, leaving just enough liquid to cover the slide.

Add 50 microliters of Gel Embedding Solution onto the tissue section.

Using tweezers, pick up the Gel Slick-treated Gel Coverslip.

With the Gel Slick-treated side facing down, place the edge of the Gel Coverslip against a second pair of tweezers resting on the slide. Slowly lower the Gel Coverslip onto the tissue section to spread the Gel Embedding Solution.

Gently press the Gel Coverslip to squeeze out excess Gel Embedding Solution.

Remove excess gel embedding solution by aspiration.

Incubate at room temperature for 1.5 hours.

Monitor the gel embedding process in the waste tube. The gel should start to form within 1 hour. Dispose of the waste tube once the gel has formed.

Gently lift the gel coverslip using the sharp tip of a blade. Discard the gel coverslip.

The tissue sample should appear very translucent.

Wash with 5 mL Sample Prep Wash Buffer. We will now proceed with tissue clearing .

Warm Clearing Premix at 37°C for 30 min before use. The Clearing Premix should be a clear solution before use. If the solution is cloudy, warm until the solution becomes clear. Prepare Clearing Solution

Add 5 milliliters of Clearing Solution. The Clearing Solution should be kept at 37 degrees C.

Cut some parafilm and use it to seal the dish.

Place dish in a larger petri dish. Cover and wipe the outside with 70% ethanol to sterilize.

Incubate in a humidified 37 degree Celsius cell culture incubator for 24 hours, or until the tissue section becomes transparent.

Once the tissue sample is completely transparent, proceed to DAPI and PolyT staining.

Prepare for Imaging
Take Sample Prep Wash Buffer and Formamide Wash Buffer from the MERSCOPE Sample Prep Kit.
Prepare a 37°C water bath with ~2 cm (height) of water in the bath (or such that the water level rises to ~2.5 cm up the outside of the cartridge). Place the applicable MERSCOPE Imaging Cartridge in the 37°C water bath for 45 min.
DO NOT allow the valve to contact the water. Warm up DAPI and PolyT Staining Reagent (PN 20300021) for 10 min in a 37°C water bath. Gently vortex the tube for 30 seconds at lowest setting to ensure the reagents are well mixed, and no precipitate is visible before use.
• Maintain Imaging Buffer Activator and RNase inhibitor in a benchtop cooler until use.
• Return unused reagents to −20°C storage but minimize freeze-thaw cycles.
• Note the MERSCOPE Imaging Cartridge barcode number in case it must be entered manually.
• Ensure the applicable panel-specific MERSCOPE Codebook is available. It may be imported from local storage or the Vizgen Cloud.
• Ensure the MERSCOPE Instrument waste container is empty before starting an experiment.

Aspirate the Clearing Solution.

Wash with 5 milliliters of Imaging Prep Wash Buffer. Do this a total of 4 times.

Add 5 milliliters of DAPI and PolyT Staining Reagent. Incubate for 10 minutes on a rocker.

Wash with 5 milliliters of Formamide Wash Buffer. Incubate for 5 minutes.

You may now proceed with image acquisition using the MERSCOPE instrument. Refer to the MERSCOPE Instrument Guide for instructions.