Jun 28, 2026
Visium HD FFPE tissue preparation and spatial gene expression protocol
- Xiang Li1,
- Feng Chen1,
- Li Ding1
- 1Washington University in St. Louis, School of Medicine
Protocol Citation: Xiang Li, Feng Chen, Li Ding 2026. Visium HD FFPE tissue preparation and spatial gene expression protocol. protocols.io https://dx.doi.org/10.17504/protocols.io.8epv5wz3nv1b/v1
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
We use this protocol and it's working
Created: June 28, 2026
Last Modified: June 28, 2026
Protocol Integer ID: 319953
Keywords: visium hd ffpe tissue preparation, complete 10x genomics visium hd spatial gene expression, spatial gene expression protocol, spatial gene expression protocol this protocol, sectioned ffpe tissue slide, visium hd slide preparation, immunofluorescence staining, rna quality assessment, probe ligation, sample index pcr
Abstract
This protocol describes the complete 10x Genomics Visium HD Spatial Gene Expression workflow, beginning with the preparation of freshly sectioned FFPE tissue slides and continuing through downstream library preparation. This version is designed for H&E staining and imaging only; immunofluorescence staining will not be performed.
The workflow includes FFPE block handling, RNA quality assessment, tissue sectioning, section placement, H&E staining and imaging, coverslip removal, destaining, decrosslinking, probe hybridization, probe ligation, Visium HD slide preparation, CytAssist-enabled probe release and capture, probe extension, probe elution, pre-amplification, SPRIselect cleanup, sample index PCR, final library cleanup, library QC, and submission.
FFPE tissue block handling for Visium HD
For best results, store FFPE tissue blocks at 4°C and protect them from direct light, high temperature, and humidity.
Before sectioning, inspect the FFPE block for tissue quality, dryness, cracks, necrosis, hemorrhage, or other obvious tissue processing artifacts.
Avoid excessive trimming, prolonged warming, or repeated freeze-thaw temperature changes, as these may affect tissue morphology and RNA quality.
Keep the tissue block chilled before sectioning. Hydrate the block on ice or a cold surface if needed, but avoid excessive exposure to water.
Clean the work area, microtome area, tools, and surrounding surfaces using RNase decontamination solution before starting.
RNA quality assessment of FFPE tissue blocks
1d
Always perform DV200 RNA quality assessment using sections from the same FFPE block that will be used for the Visium HD experiment.
Cut one or more test sections from the FFPE block for RNA extraction.
Extract RNA using an FFPE compatible RNA extraction kit.
Measure RNA concentration using nano drop and assess DV200 with TapeStation.
Calculate DV200, defined as the percentage of RNA fragments greater than 200 nucleotides.
Proceed only if RNA quality is acceptable. A DV200 greater than 30% is recommended for Visium HD.
If DV200 is low, always discuss whether to proceed or select another block, otherwise adjust expectations for downstream assay sensitivity.
Section placement on blank slides
1d
Use compatible blank slides recommended for Visium HD.
Before placing sections, mark the allowable tissue placement area (middle) on the back of the blank slide.
Hold the blank slide vertically and insert it into the water bath while aligning the allowable area with the floating tissue section.
Use a clean probe to gently guide the section into the allowable area.
Lift the slide out of the water slowly and evenly, making sure that no air bubbles are trapped underneath the tissue section.
If multiple sections are placed on one slide, ensure that tissue sections and paraffin do not overlap.
If processing multiple slides, place tissue sections in the same location on each slide whenever possible to improve imaging consistency.
Gently flick the slide to remove excess water.
Dry the slide upright at room temperature until the tissue appears opaque and no visible water remains above or underneath the section.
A fan may be used to assist drying.
Do not touch the tissue section.
Tissue slide drying and storage before H&E
3h
After section placement, dry tissue slides completely O/N before deparaffinization.
Incubate slides at 42°C for 3 hours.
If not proceeding immediately, keep tissue slides in a low moisture environment such as a desiccator.
Avoid direct light exposure.
Keep slides at 4°C unless otherwise required (can be stored for up to 6 months in desiccator).
If slides are stored at 4°C, allow them to come to room temperature before deparaffinization.
Do not use a hydrophobic barrier such as PAP pen around the tissue section.
Handle slides only with gloves.
Do not touch the tissue section.
Reagent and slide preparation for H&E workflow
20m
Prepare fresh deparaffinization reagents before starting.
Prepare two containers of xylene.
Prepare two containers of 100% ethanol.
Prepare two containers of 96% ethanol.
Prepare one container of 70% ethanol.
Prepare one container of Milli-Q.
Prepare H&E staining reagents, including hematoxylin, bluing buffer, alcoholic eosin, and water wash containers.
Prepare mounting medium. For 85% glycerol mounting medium, mix glycerol with nuclease-free water and centrifuge briefly to remove bubbles.
Prepare coverslips and lint-free laboratory wipes.
Prepare 0.1 N HCl, TE buffer pH 8.0, 1X PBS, Decrosslinking Buffer B, Perm Enzyme B, and 8 M urea for destaining and decrosslinking.
Verify Phenocycler microscope settings and imaging program before beginning staining.
Deparaffinization
3h
Retrieve tissue slides from the desiccator after storing.
If slides were stored at 4°C, place them in a slide rack at room temperature for 5 min.
Place slides tissue-side up on a Thermocycler Adapter set to 60°C and incubate for 2 h. Do not close the thermal cycler lid.
Remove slides from the thermal cycler.
Allow slides to cool to room temperature for 5 min.
Perform all xylene steps in a fume hood.
Immerse slides in Xylene Jar 1 and incubate for 10 mins.
Transfer slides to Xylene Jar 2 and incubate for 10 mins.
Move slides through the ethanol gradient using 2 x 100% ethanol, 2 x 96% ethanol, and 1 x 70% ethanol, 3 mins for each step.
Immerse slides in water for 20 secs.
Make sure all tissue sections remain fully submerged during immersion steps.
Do not allow tissue sections to dry before proceeding to H&E staining.
H&E Staining
15m
Place the slide on flat, clean, nonabsorbent work surface.
Add 1 ml hematoxylin per slide to uniformly cover all tissue sections.
Incubate for 1 min at room temperature.
Discard hematoxylin by holding the slide at an angle with the bottom edge touching a lint-free laboratory wipe.
Immerse the slide 5 times in Water Beaker 1.
Immerse the slide 15 times in Water Beaker 2.
Immerse the slide 15 times in Water Beaker 3.
Wipe excess liquid from the back of the slide without touching the tissue section.
Place the slide on a flat, clean, nonabsorbent work surface.
Add 1 ml bluing buffer per slide to uniformly cover all tissue sections.
Incubate for 1 min at room temperature.
Discard bluing buffer by holding the slide at an angle with the bottom edge touching a lint-free laboratory wipe.
Immerse the slide 15 times in Water Beaker 4.
Wipe excess liquid from the back of the slide without touching the tissue section.
Gently immerse the slide in alcoholic eosin solution in a separate 50 ml tube.
Incubate for 1 min at room temperature.
Discard eosin by holding the slide at an angle with the bottom edge touching a lint-free laboratory wipe.
Immerse the slide for 30 sec in Water Beaker 5.
Immerse the slide 10 times in Water Beaker 6.
Wipe excess liquid from the back of the slide without touching the tissue section.
Do not air dry the slide.
Coverslip Mounting
5m
Place the stained slide on a flat, clean, nonabsorbent work surface.
Some residual droplets may remain on the slide.
Using a wide-bore pipette tip, add 100-150 µl mounting medium (85% glycerol) to cover all tissue sections uniformly.
Use additional mounting medium if the tissue section is large or if multiple sections are present.
Slowly apply the coverslip at an angle from one end of the slide.
Avoid introducing bubbles.
Allow the mounting medium to spread and settle.
Do not use Cytoseal or nail polish to secure the coverslip.
Carefully wick away large excess mounting medium from the edge of the coverslip using a lint-free laboratory wipe.
Do not move the coverslip or disturb the tissue section.
Proceed immediately to imaging.
If imaging cannot be performed immediately, store slides flat at 4°C for up to 24 h.
Do not allow the coverslip to dry out.
Brightfield Imaging
2h
Remove any marker annotation with ethanol or isopropanol before imaging.
Image the tissue section using brightfield imaging settings on Phenocycler.
Use the 20x magnification and imaging format for Visium HD downstream analysis.
Ensure that the entire tissue region of interest is captured.
Check the image for focus, tissue folds, bubbles, section damage, staining artifacts, or incomplete tissue coverage.
If an area of interest is needed, identify the AOI based on the H&E image.
After imaging, proceed immediately to coverslip removal.
Coverslip Removal
15m
Dispense 800 ml Milli-Q water into a beaker.
Immerse slides sideways in the beaker, with the coverslipped surface fully sideways.
Keep the slide still in water until the coverslip slowly separates from the slide.
Do not move the slide up and down.
Do not shake the slide.
Do not manually move or force the coverslip off.
After the coverslip separates, gently immerse the slide 30 times in water to remove all mounting medium.
Wipe the back of the slide with a lint-free laboratory wipe.
Do not touch the tissue section.
Air dry the slides for 5 mins.
Incubate slides on Thermocycler adapter with lid open for 3 mins at 37°C.
Proceed immediately to destaining.
Destaining
20m
Place the Low Profile Thermocycler Adapter in the thermal cycler.
Set up the thermal cycler for destaining:
o Lid temperature: 42°C, or the lowest possible lid temperature if 42°C is not available.
o Reaction volume: 100 µl.
o Pre-equilibration: 42°C.
o Destaining temperature: 42°C.
o Destaining time: 15 min.
o Hold: 22°C.
Place the slide into a Tissue Slide Cassette.
Add 150 µl 0.1 N HCl along the side of the wells to uniformly cover the tissue sections.
Avoid bubbles.
Gently tap the cassette to ensure uniform coverage.
Remove HCl from the wells.
Add 100 µl 0.1 N HCl along the side of the wells to uniformly cover the tissue sections.
Apply a pre-cut slide seal to the cassette.
Place the cassette on the Low Profile Thermocycler Adapter at 42°C.
Close the thermal cycler lid.
Skip the pre-equilibration step and initiate destaining.
After incubation, remove the cassette from the thermal cycler and place it on a flat, clean work surface.
Peel back the slide seal.
Remove all HCl from the well corners using a pipette.
Wash 1: Add 150 µl TE buffer pH 8.0 along the side of the wells. Incubate for 5 min at room temperature. Remove TE buffer.
Wash 2: Add 150 µl TE buffer pH 8.0 along the side of the wells. Incubate for 5 min at room temperature. Remove TE buffer.
Wash 3: Add 150 µl TE buffer pH 8.0 along the side of the wells. Incubate for 5 min at room temperature. Remove TE buffer.
Add 100 µl 1X PBS along the side of the wells.
Re-apply the slide seal.
Proceed directly to decrosslinking.
Decrosslinking
40m
Thaw Decrosslinking Buffer B in thermomixer (300 rpm shaking) for 30 min at 37°C before starting, then cool to room temperature for 5 min. Vortex for 30 sec and centrifuge briefly.
Equilibrate Perm Enzyme B to room temperature shortly before use, pipette mix and briefly centrifuge.
Set up the thermal cycler for decrosslinking:
o Lid temperature: 80°C.
o Reaction volume: 100 µl.
o Hold: 22°C.
o Decrosslinking: 80°C for 30 min.
o Re-equilibration: 22°C for 10 min.
o Hold: 22°C.
Prepare Diluted Perm Enzyme B:
o 998 µl 1X PBS.
o 2 µl Perm Enzyme B.
o Mix thoroughly with a 1 ml pipette set to 600 µl.
o Maintain at room temperature.
Prepare Decrosslinking Mix (per slide):
o 92.5 µl Decrosslinking Buffer B.
o 6.25 µl 8 M urea.
o 1.25 µl Diluted Perm Enzyme B.
o Total volume: 100 µl.
Pipette mix thoroughly and briefly centrifuge, then maintain at room temperature.
Remove the slide seal.
Remove 1X PBS from the wells.
Add 100 µl Decrosslinking Mix along the side of the wells.
Apply a new pre-cut slide seal to the cassette.
Place the cassette on the Low Profile Thermocycler Adapter.
Close and tighten the thermal cycler lid.
Skip the pre-equilibration step and initiate decrosslinking.
After decrosslinking is complete, proceed immediately to the Visium HD Spatial Gene Expression Reagent Kits User Guide.
Probe hybridization
1d
Prepare Pre-Hybridization Mix
Prepare Pre-Hybridization Mix shortly before use.
Add reagents in the following order per slide:
o Nuclease-free water: 134.2 µl.
o 10X PBS, pH 7.4: 15 µl.
o 10% Tween-20: 0.8 µl.
For each 6.5 mm sample, prepare 150 µl Pre-Hybridization Mix.
Vortex the mix thoroughly.
Centrifuge briefly.
Keep Pre-Hybridization Mix at room temperature.
Remember to thaw FFPE Hyb Buffer at 37°C until dissolved, pipette mix 10x then keep it at room temperature.
Remember to thaw Human or Mouse WT Probes – RHS/LHS at room temperature, vortex and centrifuge briefly, maintain at room temperature.
Add Pre-Hybridization Mix to tissue slides
Retrieve tissue slide cassettes containing H&E-stained, destained, and decrosslinked tissue sections.
Place each tissue slide cassette flat on a clean work surface.
Carefully peel back the Visium Slide Seal.
Remove all buffer from each well using a pipette.
Use a P20 pipette to remove any remaining liquid from the well corners.
Add 150 µl Pre-Hybridization Mix along the side of each well.
Make sure the tissue section is uniformly covered.
Avoid introducing bubbles.
Re-apply the Visium slide seal.
Incubate for 15 min at room temperature.
Prepare thermal cycler for probe hybridization
Place a Low Profile Thermocycler Adapter in the thermal cycler.
Set up the thermal cycler using the following program:
o Lid temperature: 50 °C
o Reaction volume: 100 µl
o Pre-equilibrate: 50°C hold
o Hybridization: 50°C for 16-24 h
o Post-hybridization wash: 50°C hold
Start the program and allow the adapter to equilibrate.
Prepare Probe Hybridization Mix
Prepare Probe Hybridization Mix shortly before use.
Add reagents in the following order:
o Nuclease-free water: 10 µl.
o FFPE Hyb Buffer: 70 µl.
o Human WT Probes v2 RHS or Mouse WT Probes v2 RHS: 10 µl.
o Human WT Probes v2 LHS or Mouse WT Probes v2 LHS: 10 µl.
For each 6.5 mm sample, prepare 100 µl Probe Hybridization Mix.
Pipette mix 10 times.
Centrifuge briefly.
Keep Probe Hybridization Mix at room temperature.
Add Probe Hybridization Mix
Peel back the Visium slide seal.
Remove all Pre-Hybridization Mix from each well.
Add 100 µl Probe Hybridization Mix to each well.
Add reagent along the side of the well.
Ensure the tissue is completely covered.
Avoid bubbles.
Apply a new pre-cut Visium slide seal to each tissue slide cassette.
Place the Tissue Slide Cassette on the Low Profile Thermocycler Adapter.
Close the thermal cycler lid.
Skip the pre-equilibration step and start the hybridization step.
Incubate at 50°C overnight for 16-24
Probe ligation
1h 55m
Post-hybridization wash
25m
Prepare wash buffers
Thaw FFPE Post-Hyb Wash Buffer at 37°C until dissolved, vortex and centrifuge briefly.
Thaw 2X Probe Ligation Buffer at room temperature until no precipitate remains, vortex and centrifuge briefly.
Aliquot FFPE Post-Hyb Wash Buffer (495 µl FFPE Post-Hyb Wash Buffer per slide).
Pre-heat FFPE Post-Hyb Wash Buffer to 50°C.
Maintain the buffer at 50°C throughout the wash steps.
Prepare 2X SSC Buffer (1500 µl per slide).
Keep 2X SSC Buffer at room temperature.
Do not discard excess 2X SSC Buffer, as it will be used again in later steps.
Perform post-hybridization washes
Remove tissue slide cassettes from the thermal cycler.
Place cassettes flat on a clean work surface.
Carefully peel back the Visium slide seal.
Remove all Probe Hybridization Mix from each well.
Immediately add 150 µl pre-heated FFPE Post-Hyb Wash Buffer to each well.
Re-apply the Visium slide seal.
Place the cassette back on the pre-heated Low Profile Thermocycler Adapter.
Incubate at 50°C for 5 min.
Remove the cassette from the thermal cycler.
Peel back the Visium slide seal.
Remove all FFPE Post-Hyb Wash Buffer from each well.
Immediately add 150 µl pre-heated FFPE Post-Hyb Wash Buffer.
Re-apply the Visium slide seal.
Incubate at 50°C for 5 min.
Repeat the wash one more time for a total of three post-hybridization washes.
After the final wash, remove all FFPE Post-Hyb Wash Buffer.
Add 150 µl 2X SSC Buffer to each well.
Re-apply the Visium slide seal.
Allow the tissue slide cassettes to cool to room temperature for ~3 min.
Probe ligation
1h 5m
Prepare thermal cycler
Place the Low Profile Thermocycler Adapter on the thermal cycler.
Set up the thermal cycler using the following program:
o Lid temperature: 37°C
o Reaction volume: 100 µl
o Pre-equilibrate: 37°C hold
o Ligation: 37°C for 1 h
o Hold: 4°C
Start the program and allow the adapter to equilibrate.
Prepare Probe Ligation Mix
Prepare Probe Ligation Mix shortly before use.
2X Probe Ligation Buffer is pre thawed. Also get Probe Ligation Enzyme shortly, centrifuge briefly and maintain on ice.
Add reagents in the following order per slide:
o Nuclease-free water: 24 µl.
o 2X Probe Ligation Buffer: 30 µl.
o Probe Ligation Enzyme: 6 µl.
For each 6.5 mm sample, prepare 60 µl Probe Ligation Mix.
Pipette mix 10 times.
Centrifuge briefly.
Do not vortex.
Keep Probe Ligation Mix on ice.
Perform probe ligation
Remove the Visium slide seal from each tissue slide cassette.
Remove all 2X SSC Buffer from each well.
Add 60 µl Probe Ligation Mix directly to the tissue section in each well.
Avoid introducing bubbles.
Gently tap the cassette to ensure uniform reagent coverage.
Apply a new pre-cut Visium slide seal.
Place the cassette on the pre-heated Low Profile Thermocycler Adapter.
Close the thermal cycler lid.
Skip the pre-equilibration step.
Start ligation.
Incubate at 37°C for 1 h.
Post-ligation wash
25m
Prepare Post-Ligation Wash Buffer
Thaw Post-Ligation Wash Buffer at room temperature right after ligation incubation.
If proceeding directly to Visium HD Slide Preparation, remove the Visium HD slide mailer from -80°C and thaw at room temperature for 30-60 min.
Keep the slide mailer upright and capped.
Pre-heat Post-Ligation Wash Buffer to 57°C shortly before use.
Use 110 µl Post-Ligation Wash Buffer per 6.5 mm sample.
First post-ligation wash
Set the thermal cycler to 57°C.
After ligation, remove tissue slide cassettes from the thermal cycler.
Place cassettes flat on a clean work surface.
Remove the Visium slide seal.
Remove all Probe Ligation Mix from each well.
Immediately add 100 µl room temperature Post-Ligation Wash Buffer to each well.
Apply a new pre-cut Visium slide seal.
Place the cassette on the Low Profile Thermocycler Adapter.
Close the thermal cycler lid.
Incubate at 57°C for 5 min.
Second post-ligation Wash
Remove the cassette from the thermal cycler.
Place it on a clean work surface.
Peel back the Visium slide seal.
Remove all Post-Ligation Wash Buffer from each well.
Add 100 µl pre-heated Post-Ligation Wash Buffer to each well.
Re-apply the Visium slide seal.
Place the cassette back on the Low Profile Thermocycler Adapter.
Incubate at 57°C for 5 min.
Final 2X SSC wash
Remove the cassette from the thermal cycler.
Peel back the Visium slide seal.
Remove all Post-Ligation Wash Buffer from each well.
Add 150 µl 2X SSC Buffer to each well.
Remove all 2X SSC Buffer from each well.
Add fresh 150 µl 2X SSC Buffer to each well.
Re-apply the Visium Slide Seal.
Allow tissue slide cassettes to come to room temperature for 5 min.
Store at 4°C for up to 24 h or proceed directly to Visium HD Slide preparation.
Visium HD Slide preparation
25m
Visium HD Slide wash
Prepare Visium HD slide
Confirm that the Visium HD slide mailer has thawed at room temperature for 30-60 min.
Keep the slide mailer upright and capped during thawing.
Prepare 60 ml 0.1X SSC Buffer.
Prepare a new 6.5 mm Visium cassette.
Record the Visium HD slide serial number before placing the slide into the cassette.
Handle the Visium HD slide gently.
Ensure the active surface is facing up.
Do not touch the active surface.
Wash and equilibrate the Visium HD Slide
Open the slide mailer carefully.
Wash the slide using 7 ml 0.1X SSC Buffer slowly along the side of mailer on one side of the slide.
Wash the slide using 7 ml 0.1X SSC Buffer slowly along the side of mailer on another side of the slide, so both side of slide are immersed with buffer.
Incubate at room temperature for 1 min.
Remove the 0.1X SSC Buffer.
Wash the slide using 7 ml 0.1X SSC Buffer slowly along the side of mailer on one side of the slide.
Wash the slide using 7 ml 0.1X SSC Buffer slowly along the side of mailer on another side of the slide, so both side of slide are immersed with buffer.
Incubate at room temperature for 5 min.
Remove the 0.1X SSC Buffer.
Repeat two more washes for a total of three 5 min washes.
Ensure back and side of Visium HD slide is dry by using lint-free lab wipe.
Ensure Visium HD slide is free of dust, if necessary rewash the slide with 0.1X SSC Buffer.
Place the Visium HD slide into a new 6.5 mm Visium Cassette.
Add 100 µl 0.1X SSC Buffer to each well of the cassette.
Apply a new pre-cut Visium Slide Seal.
Leave the Visium HD slide in 0.1X SSC Buffer at room temperature until the equilibration step.
Do not exceed 2 h before proceeding to the CytAssist run.
Keep the Visium Cassette and Visium HD slide free from dust and debris.
Probe release and extension
2h 55m
CytAssist-enabled probe release and capture
1h 10m
Prepare CytAssist run
Confirm that the CytAssist firmware version is compatible with the Visium HD workflow.
Prepare the CytAssist instrument.
Record which tissue slide corresponds to each Visium HD Capture Area.
Record whether each Capture Area is on the left or right side of the Visium Slide Stage.
Confirm that each tissue section fits within the 6.5 mm alignment guide area.
If only one tissue slide is processed, load a blank slide in the second position.
Do not reuse the unused Capture Area for another CytAssist run.
Reagent preparation
Prepare 6600 µl 1x PBS per two slides.
Prepare 330 µl 10% Eosin per two slides:
o Alcoholic Eosin: 33 µl.
o 1X PBS: 297 µl.
For preparation of Visium HD slide Equilibration Mix, firstly thaw 2x RNase Buffer at room temperature, pipette mix slowly, keep at room temperature. Then get RNase Enzyme shortly before use, pipette mix and centrifuge briefly, keep on ice.
Equilibration and loading the Visium HD Slide for CytAssist
Prepare Slide Equilibration Mix per HD slide:
o Nuclease-free Water: 96.8 µl.
o 2x RNase Buffer: 11 µl.
o RNase Enzyme: 13.2 µl.
o Total: 220 µl.
Remove Visium slide seal and 0.1X SSC Buffer from the Visium cassette wells for HD slide.
Add 100 µl HD Slide Equilibration Mix to each well in the cassette, gently tap to ensure uniform coverage the capture area.
Apply a new uncut Visium slide seal on the Visium cassette and incubate at room temperature for 10 min.
Prepare Probe Release Mix during the equilibration per two tissue slides:
o 2x RNase Buffer: 20 µl.
o RNase Enzyme: 17.5 µl.
o Total: 37.5 µl.
Remove Slide Equilibration Mix completely from the Visium cassette wells for HD slide.
Remove the Visium HD slide from the cassette.
Load the Visium HD slide onto the Visium Slide Stage.
Make sure the label is facing up and oriented correctly.
Fit the slide into the raised grooves.
Close the Visium Slide Lock gently.
Confirm the Visium HD slide is seated correctly.
Allow the Visium HD slide to dry on the Visium Slide Stage for 10 min.
Inspect the entire well area.
Continue drying if any liquid remains.
Prepare tissue slides
Retrieve the tissue slide cassettes.
Remove all 2X SSC Buffer from each well.
Remove tissue slides from the Tissue Slide Cassettes.
Apply 100 µl 10% eosin to uniformly cover each tissue section.
Incubate 1 min at room temperature.
Remove eosin.
Rinse the tissue with 1 ml 1x PBS, do not rinse directly onto tissue.
Repeat two more times for a total of three 1x PBS washes.
Dry the tissue slide completely by waft the tissue slides.
Wipe back of tissue slides with lint-free lab wipe.
Inspect the entire tissue area and well area to confirm no liquid remains.
Do not proceed if visible liquid remains.
Load tissue slides onto CytAssist
Open the tissue slide clips.
Place the tissue slide flat on the Tissue Slide Stage.
Align the tissue section or area of interest within the 6.5 mm alignment guides.
Position high-priority tissue regions inside the alignment guide area.
If needed, rotate the tissue slide 180° to improve tissue placement.
Secure the slide using the clips.
Adjust the slide gently until the tissue is centered within the guide area.
Confirm the slide does not overlap the center line.
Load a second tissue slide or a blank slide into the second position.
Confirm both slides are secured.
Add Probe Release Mix and run CytAssist
Get Perm Enzyme B shortly before use, pipette mix, centrifuge briefly and maintain at room temperature.
Add 2.5 µl of Perm Enzyme B to 37.5 µl of Probe Release Mix prepared in 4.1.3, pipette mix 15x wit pipette set to 30 µl without generating bubbles, centrifuge for 5 sec.
The time between adding Perm Enzyme B to Probe Release Mix and starting Visium CytAssist run should be less than 5 min.
Slowly add 17 µl of Probe Release Mix into the center of each spacer well on the Visium HD slides, avoid generating bubbles.
Close lid slowly and start the CytAssist run ~30 mins.
Probe extension
1h 25m
Prepare incubation protocol for probe extension
During CytAssist run, set up the thermal cycler using the following program:
o Lid temperature: 53°C
o Reaction volume: 100 µl
o Pre-equilibrate: 53°C hold
o Probe Extension 1: 53°C for 30 mins o Cool Down: 4°C for 3 mins
o Hold : 4°C
o Probe Extension 2: 53°C for 30 mins
o Cool Down: 4°C for 3 mins
o Hold : 4°C
Prepare Probe Extension Mix
During CytAssist run, thaw Extension Buffer at room temperature, then vortex and centrifuge briefly, keep at room temperature.
Get Extension Enzyme out shortly before using, pipette mix, centrifuge briefly and keep on ice.
Prepare Probe Extension Mix per slide.
o Extension Buffer: 323.4 µl.
o Extension Enzyme: 6.6 µl.
o Total: 330 µl.
Pipette mix, centrifuge briefly and keep Probe Extension Mix on ice.
Probe extension round 1
After the CytAssist run is complete, immediately remove the Visium HD slide.
It is normal if tissue remains on the tissue slide after the CytAssist run.
Hold the Visium HD slide over a liquid waste container.
Rinse each capture area with 1 ml Buffer EB.
Do not pipette directly onto the Capture Area.
Repeat the Buffer EB wash two more times for a total of three washes per capture area.
Place the Visium HD slide back into the same Visium cassette.
Proceed immediately to Probe Extension.
Add 75 µl Probe Extension Mix to each well of the Visium cassette.
Gently tap the cassette to ensure uniform coverage of each capture area.
Keep the remaining Probe Extension Mix on ice.
Apply a new uncut Visium Slide Seal.
Place the Visium Cassette on the pre-heated Low Profile Thermocycler Adapter.
Close the thermal cycler lid.
Skip the pre-equilibration step.
Start Probe Extension 1.
Probe extension round 2
After the first cool down step, open the thermal cycler lid.
Remove the Visium cassette from the Low Profile Thermocycler Adapter.
Peel back the Visium Slide Seal.
Remove Probe Extension Mix from each well.
Add 75 µl fresh Probe Extension Mix to each well.
Gently tap the cassette to ensure uniform coverage.
Re-apply the Visium Slide Seal.
Place the cassette back on the Low Profile Thermocycler Adapter.
Close the thermal cycler lid.
Skip the hold step and start Probe Extension 2.
After extension, samples may remain at 4°C in the thermal cycler for up to 24 h.
Probe elution
20m
Prepare probe elution mix per slide.
o Nuclease-free Water: 108.9 µl.
o KOH (8 M): 1.1 µl.
o Total: 110 µl.
Remove the Visium cassette from the thermal cycler.
Place it on a clean, flat work surface.
Peel back the Visium slide seal.
Remove all Probe Extension Mix from each well.
Add the 50 µl elution buffer to each well.
Gently tap the cassette to ensure uniform coverage.
Incubate at room temperature for 10 min.
During incubation, prepare two PCR tubes, and add 3 µl 1 M Tris-HCl (pH 8.0) to the tube for each sample.
Collect the eluate from each well of Visium cassette completely to the PCR tubes containing Tris-HCl, record the sample information.
Vortex, centrifuge briefly and keep samples on ice.
Proceed immediately to Pre-Amplification.
Pre-amplification and SPRIselect cleanup
50m
Pre-amplification
Prepare Pre-Amplification Mix
During probe elution step, thaw TS Primer Mix B at room temperature, vortex and centrifuge briefly, keep at room temperature.
Get Amp Mix B shortly before using, vortex, centrifuge briefly and keep on ice.
Prepare Pre-Amplification Mix on ice.
Add reagents in the following order per sample:
o Nuclease-free water: 42.9 µl.
o Amp Mix B: 55.0 µl.
o TS Primer Mix B: 5.5 µl.
o Total: 103.4 µl.
Pipette mix, centrifuge briefly and maintain on ice.
Set pre-amplification PCR
Use the following thermal cycler program:
o Lid temperature: 105°C
o Reaction volume: 100 µl
o 98°C for 3 min
o 98°C for 15 sec
o 63°C for 20 sec
o 72°C for 30 sec
o Repeat steps 98°C (15 sec)-72°C (30 sec) for a total of 10 cycles
o 72°C for 1 min
o Hold at 4°C
Run pre-amplification reaction
Add 47 µl Pre-Amplification Mix to each PCR tube from probe elution.
Pipette mix and centrifuge briefly.
Place reactions in a thermal cycler and start the run.
After PCR, proceed directly to SPRIselect cleanup.
Pre-amplification cleanup - SPRIselect
Vortex SPRIselect reagent (~30 sec) to fully resuspend beads.
Add 120 µl SPRIselect beads to each 100 µl pre-amplification sample.
Pipette mix 15 times with pipette set to 175 µl.
Incubate for 5 min at room temperature.
Place the tube strip on the magnet with high pattern until the solution clears (~3 min).
Remove and discard the supernatant.
Add 300 µl freshly prepared 80% ethanol to the bead pellet.
Wait 30 sec.
Remove ethanol.
Add 200 µl freshly prepared 80% ethanol to the bead pellet.
Wait 30 sec.
Remove ethanol.
Briefly centrifuge the tube strip.
Place the tube strip back on the magnet with low pattern.
Remove any remaining ethanol without disturbing the beads.
Remove tube strip from the magnet.
Add 105 µl Buffer EB.
Pipette mix 15 times with pipette set to 100 µl.
Incubate for 2 min at room temperature.
Place the tube strip on the magnet with high pattern until the solution clears (~ 3 mins).
Transfer 100 µl cleaned sample to a new tube strip.
Store at 4°C for up to 72 h, at -20°C for up to 4 weeks, or proceed directly to library construction.
Visium HD probe-based library construction
2h 35m
Cycle number determination by qPCR
1h
Prepare diluted TS Primer Mix B
Thaw TS Primer Mix B at room temperature, vortex, centrifuge briefly and keep at room temperature.
Dilute TS Primer Mix B 1:10 in nuclease-free water.
Vortex and centrifuge briefly.
Keep diluted primer mix on ice.
Prepare qPCR Mix
Prepare qPCR Mix on ice.
Add reagents in the following order per well:
o KAPA SYBR FAST qPCR Master Mix: 5 µl.
o Diluted TS Primer Mix B: 1 µl.
o Nuclease-free water: 3 µl.
o Total: 9 µl.
Minimize light exposure.
Vortex the qPCR Mix, centrifuge briefly and maintain on ice.
Set up qPCR reaction
Add 9 µl qPCR Mix to one qPCR plate well per sample.
Include a negative control well.
Dilute 2 µl cleaned pre-amplified sample in 8 µl nuclease-free water.
Pipette mix and centrifuge briefly.
Transfer 1 µl diluted sample into the corresponding qPCR well.
For the negative control, add 1 µl nuclease-free water.
Pipette mix the qPCR each well.
Seal the qPCR plate.
Centrifuge briefly.
Record sample positions in the qPCR plate.
Run qPCR and Determine Cycle Number
Run qPCR using following qPCR program:
o Lid temperature: 105°C
o Reaction volume: 10 µl
o 98°C for 3 min
o 98°C for 5 sec
o 63°C for 30 sec
o Read signal
o Repeat steps 98°C (5 sec)-Read signal for a total of 25 cycles
Review amplification curves and record the Cq value (~25% of the peak fluorescence value) for each sample.
Determine the appropriate sample index PCR cycle number based on the Cq (Cq + 1) result.
Record the selected cycle number for each sample.
Sample index PCR
40m
Prepare sample index PCR reaction
Thaw Dual Index Plate TS Set A during qPCR run.
Centrifuge the index plate briefly.
Choose an appropriate unique index well for each sample.
Record the index well used for each sample.
Use each index well only once to avoid index overlap in multiplexed sequencing runs.
Prepare Amplification Master Mix per sample, pipette mix 10x shortly before use:
o Nuclease-free Water: 45 µl.
o Amp Mix B: 25 µl.
o Total: 70 µl.
Add the 70 µl of Amplification Master Mix, then add 25 µl of cleaned pre-amplified sample into a new PCR tube.
Add the 5 µl selected sample index from Dual Index TS Set A.
Mix thoroughly and centrifuge briefly.
Run sample index PCR
Set sample index PCR using following program:
o Lid temperature: 105°C
o Reaction volume: 100 µl
o 98°C for 3 min
o 98°C for 15 sec
o 63°C for 20 sec
o 72°C for 30 sec
o Repeat steps 98°C (15 sec)-72°C (30 sec) for a total of Cq + 1 cycles
o 72°C for 1 min
o 4°C hold
Place reactions in the thermal cycler.
Run the sample index PCR program using the cycle number determined by qPCR.
After PCR, keep samples on ice.
Proceed directly to post-sample index PCR cleanup.
Post-sample index PCR cleanup - SPRIselect
25m
Vortex SPRIselect reagent (~30 sec) to fully resuspend beads.
Add 85 µl of SPRIselect reagent to each Sample Index PCR sample.
Pipette mix 15 times with pipette set to 175 µl.
Incubate 5 mins at room temperature.
Place the tube strip on the magnet at high pattern until the solution clears (~ 3 min).
Remove and discard the supernatant.
Wash beads with 200 µl freshly prepared 80% ethanol.
Remove ethanol.
Repeat the ethanol wash for a total of two washes.
Place the tube strip on the magnet at low pattern.
Remove residual ethanol completely.
Do not over-dry beads.
Remove tube strip from the magnet.
Elute DNA using 27 µl Buffer EB.
Pipette mix 15 times.
Incubate 2 mins at room temperature.
Place tube strip back on the magnet at low pattern until the solution clears (~ 3 min).
Transfer 25 µl cleaned final library to a new PCR tube.
Store final library at -20°C for long-term storage.
Post-library construction QC
30m
Assess library size distribution using TapeStation using diluted samples (1:50 dilution).
Confirm that final library traces show the expected size distribution (the expected average fragment size is 250 bp).
Check for abnormal traces, such as low yield, flat trace, overamplification, adaptor dimers, or overloaded peaks.
Record library concentration.
Record average library size.
Dilute libraries to the required loading concentration for sequencing.
Pool libraries according to sequencing plan.
Store libraries at -20°C if not sequencing immediately.