Agarose-Assisted Paraffin Embedding & H&E Staining Guide for Organoids

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July 09, 2026

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The embedding workflow of organoids is basically consistent with that of bulk tissues. However, organoids exist as numerous discrete microstructures, whereas tissues are integrated bulk aggregates. Organoids are prone to dispersion and loss during dehydration. To facilitate handling during embedding and precisely locate sectioning regions, agarose-assisted embedding is required. Organoids vary in morphology and culture protocols depending on their tissue origin, leading to distinct collection difficulties. Two distinct agarose embedding pre-treatment methods prior to dehydration can be selected according to organoid characteristics.

Conventional Agarose Embedding Method: Applicable to suspension-cultured organoids, vacuolar organoids that remain intact after matrix removal, and compact solid organoids.

In-situ Agarose Embedding Method: Applicable to fragile organoids prone to fragmentation post matrix digestion, or samples with low organoid yield.

I. Organoid Agarose Embedding

1. Conventional Agarose Embedding Protocol

(1) Organoid Harvesting

a. Harvest organoids when their diameter reaches approximately 200 μm.

b. Aspirate culture medium with a pipette, add 1–2 mL pre-chilled (4°C) Organoid Passaging Buffer (abs9730) to each well and incubate for 2 min with gentle manipulation throughout the procedure.

c. Gently triturate Matrigel droplets, collect suspensions into 15 mL centrifuge tubes, and statically incubate at 4°C for 30 min (group every 4 wells).

d. Centrifuge for 5 min and discard supernatant. Remove residual Matrigel using a 200 μL pipette; maximize residual gel elimination. Resuspend pelleted organoids in tissue fixative, then transfer to 1.5 mL EP tubes (Pre-rinse pipette tips with serum to prevent organoid adhesion to tube walls during transfer).

(2) Agarose Embedding Procedure

a. Weigh 0.2–0.4 g agarose powder and dissolve in 10 mL deionized water buffer. Heat in a water bath at 65–70°C until fully melted. Transfer 0.6 mL liquefied agarose solution into a new 1.5 mL centrifuge tube via pipette.

b. Insert a 0.5 mL centrifuge tube into the 1.5 mL tube filled with agarose. After agarose solidifies, extract the 0.5 mL tube to form a recessed cavity.

c. Pre-stain fixed organoids with 30 μL Eosin Y, mix by gentle inversion, then centrifuge at 1000 rpm for 5 min. Remove most supernatant while retaining 20–30 μL liquid above the pellet; resuspend the pellet thoroughly and transfer suspension into the agarose cavity prepared in step b. Overlay the agarose surface slightly with PBS buffer.

d. Centrifuge at 1000 rpm for 5 min and discard supernatant. Repeat this step twice to eliminate residual paraformaldehyde and Eosin Y.

e. After centrifugation, add adequate molten agarose into the cavity and gently triturate with a pipette to fully encapsulate organoids within agarose matrix.

f. Allow agarose to solidify at room temperature, then extract the agarose block.

g. Place agarose blocks into centrifuge tubes and incubate in paraformaldehyde fixative for 12 h.

2. In-situ Agarose Embedding Protocol

(1) Organoid Harvesting

a. Harvest organoids when their diameter reaches approximately 200 μm.

b. Aspirate culture medium carefully (Avoid contact with Matrigel droplets), add 1.5 mL paraformaldehyde directly into each well for 12 h fixation.

(2) Agarose Embedding Procedure

a. Add 20 μL Eosin Y into each well of a 24-well plate, gently triturate to homogenize, and incubate statically for 5 min to pre-stain Matrigel droplets.

b. Gently detach Matrigel droplets from the well bottom using a pipette tip. Discard paraformaldehyde and wash thoroughly with PBS to remove residual fixative.

c. Dissolve 0.2–0.4 g agarose powder in 10 mL deionized water buffer, heat at 65–70°C in water bath until complete melting. Pipette molten agarose into each 24-well plate well, then adjust droplet position with pipette tips to suspend Matrigel droplets within agarose solution.

d. After cooling and solidification, trim excess agarose to retain only the matrix region containing Matrigel droplets.

After mastering the two agarose embedding protocols above, we proceed to agarose block dehydration and paraffin embedding procedures.

II. Dehydration

1. Gradient ethanol serial dehydration: Immerse samples in 70%, 80%, 90% and 95% ethanol for 40 min each sequentially.

2. Immerse in absolute ethanol for 40 min, repeat twice for complete dehydration.

3. Incubate samples in 1:1 mixture of absolute ethanol and xylene for 30 min.

4. Clear in xylene twice, 30 min per incubation.

5. Post clearing, transfer samples to wax bath at 60°C for paraffin infiltration, repeat twice with 30 min per cycle, followed by paraffin embedding.

III. Paraffin Embedding

1. Dispense molten paraffin into embedding molds, place agarose blocks at the mold center, then transfer molds to the cryostat cooling platform. Once the bottom paraffin turns opaque and solidifies, mount embedding cassettes onto molds and fill cassettes with molten paraffin.

2. Keep embedding molds on cooling platform until paraffin fully solidified prior to demolding.

IV. Sectioning

1. Cool paraffin blocks at -10°C for 5 min, then clamp blocks onto microtome specimen holders.

2. Adjust blade holder-to-block distance for rough trimming until approximately 80% of the agarose block surface is exposed.

3. Perform fine trimming until roughly 95% of agarose matrix is exposed, or organoid cells are visible under light microscopy.

4. Float trimmed sections onto 45°C warm water bath for flattening until wrinkle-free, then retrieve sections with glass slides.

5. Adjust section thickness appropriately according to tissue and agarose block properties.

6. Label slides with unique specimen IDs, bake slides in oven at 65°C for 2 h.

V. Hematoxylin-Eosin (H&E) Staining Protocol

1. Deparaffinization & Rehydration

(1) Deparaffinization: Sequentially immerse paraffin sections in three separate xylene baths for 5 min each to dissolve paraffin wax.

(2) Rehydration: Serial gradient ethanol from 100% down to 70%, 5 min per concentration. Final rinse with distilled water for 1 min; slides without water haze are ready for downstream staining.

2. Hematoxylin Nuclear Staining

(1) Staining: Cover sections with hematoxylin working solution for 5 min, then rinse under running tap water to remove excess stain.

(2) Differentiation: Treat slides with differentiation solution for 10 sec, rinse thoroughly under running tap water to eliminate residual differentiator.

(3) Blueing: Incubate slides in bluing reagent for 10 sec, rinse under running tap water to wash off surplus bluing solution.

3. Eosin Cytoplasmic Counterstaining

Staining procedure: Cover slides with Eosin Y working solution for 2 min, rinse under running tap water to remove excess Eosin.

4. Dehydration & Clearing

(1) Dehydration: Serial gradient ethanol from 70% up to 100%, 2 min per concentration for complete dehydration.

(2) Clearing: Transfer slides into two independent xylene baths, incubate 5 min each to render tissue transparent.

5. Mounting

Mounting procedure: Dispense adequate neutral mounting medium onto tissue sections, apply coverslips slowly to avoid air bubble formation, then air-dry slides naturally.

This tutorial concludes here. Feel free to join our technical communication group for experimental troubleshooting!

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