Method Article

Mapping Hepatic Stellate Cell Morphology in Mouse Models of Liver Fibrosis

DOI:

10.3791/70396

⸱

February 13th, 2026

 , 
1Division of Liver Diseases, Icahn School of Medicine at Mount Sinai

In This Article

Summary

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This protocol presents a comprehensive imaging pipeline to visualize and model hepatic stellate cells morphology in intact liver tissue. This approach combines fluorescent labeling of fibroblasts using mouse genetics, in situ liver perfusion-based tissue preparation, and a modified iDISCO tissue clearing method to achieve optical transparency.

Abstract

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Tissue-resident fibroblasts are the cellular drivers of fibrosis across tissues. Within the liver, hepatic stellate cells (HSCs), the resident pericytes with their unique "star-like" morphology, activate into fibrogenic myofibroblasts during injury to produce the hepatic scar. While decades of work elucidated major signaling pathways underlying HSC activation, the functional significance of their distinct dendritic morphology remains elusive due to technical limitations in imaging and analysis. This protocol presents a comprehensive imaging pipeline to visualize and model HSC morphology in intact liver tissue. This approach combines fluorescent labeling of HSCs using mouse genetics, in situ liver perfusion-based tissue preparation, and a modified iDISCO tissue clearing method to achieve optical transparency in tissue specimens that can be several centimeters thick. Cleared samples are imaged using optimized confocal microscopy platforms, and image datasets are processed through a customized analysis workflow for three-dimensional reconstruction of individual HSCs at single-cell resolution. Together, this pipeline enables reproducible mapping of HSC morphology within the complex liver microenvironment, including their dendritic processes and spatial relationships to neighboring cells. Beyond the liver, the workflow is adaptable to fibroblasts in other organs, such as the lung, kidney, intestine, and heart, where resident mesenchymal cells exhibit similar fibrogenic roles in disease. The proposed framework, therefore, establishes a methodological foundation for comparative cross-organ studies to probe fibroblast structure-function relationships and therapeutic vulnerabilities in fibrotic disease.

Introduction

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Fibrosis, or the pathological scarring of tissue, is the final common pathway for nearly all chronic organ diseases and contributes to approximately 40% of deaths in industrialized nations1. In the liver, fibrosis arises from chronic insults such as viral hepatitis, alcohol use, or increasingly, metabolic dysfunction-associated steatotic liver disease (MASLD), which currently affects 30-40% of adults in the U.S. Chronic insults to the liver and the resulting progressive fibrosis drastically increase the risk of end-stage liver diseases such as cirrhosis, liver failure, and hepatocellular carcinoma2,3,4,5.

Hepatic stellate cells (HSCs) are central to liver homeostasis and fibrogenesis. In healthy livers, HSCs exist in a quiescent, vitamin A-storing state and contribute to the homeostasis of hepatocytes and, overall, the entire liver6,7. This is clearly demonstrated by HSC depletion studies, where the removal of nearly all HSCs in healthy liver decreases liver mass and severely limits liver regeneration7,8. Upon injury, HSCs activate into myofibroblast-like cells that proliferate, migrate to damaged areas, and produce excessive extracellular matrix (ECM), forming fibrotic scars. Activated HSCs in fibrotic livers are also the source of cancer-associated fibroblasts and play tumor-promoting roles in hepatocellular carcinoma9,10,11,12. Quiescent HSCs possess a unique neuron-like morphology with small cell bodies from which long-range projections extend. HSCs shift to a flat, sheet-like myofibroblast shape when activated in fibrosis13. While decades of work have elucidated the signaling pathways controlling ECM production by HSCs, the functional significance of their distinct neuronal morphology, how it is regulated, whether it could facilitate interactions within the HSC niche, and contribute to HSC fibrogenicity and disease remain elusive14,15.

Current methods to visualize HSCs' morphology are limited to two-dimensional (2D) images of either in vitro cultured cells or in vivo tissue sections, and rarely capture the entirety of HSC morphology with projections extending several cell body lengths in all directions. Moreover, the most commonly used HSC marker, desmin, cannot fully capture the unique neuron-like morphology of the HSCs in immunofluorescence staining, as desmin is a cytoskeleton protein that only labels intermediate filaments within HSCs16. These methodological challenges thus far hindered the ability to fully capture HSC morphology and their cellular interactions in a three-dimensional (3D) manner in their natural context at single-cell resolution. As a result, fundamental questions regarding how HSC morphology relates to their functional state and interactions with neighboring cell types have remained unresolved.

This article presents an innovative and comprehensive imaging and analysis pipeline to visualize and model HSC morphology in intact liver tissue. This method integrates several key innovations: (1) fluorescent labeling of HSCs using genetic reporter mouse models, (2) optimized in situ liver perfusion and fixation to preserve intact tissue architecture and cellular relations, (3) a modified iDISCO-based tissue clearing protocol to achieve optical transparency17, (4) high-resolution confocal microscopy for deep-tissue image capture, and (5) a customized computational workflow for 3D image reconstruction and quantitative analysis of individual HSCs. The protocol imposes certain constraints on tissue size and time commitment that need to be taken into consideration. Together, this protocol provides a robust and reproducible framework for capturing the complex 3D structure of HSCs and for mapping their spatial relationships with neighboring vascular, immune, and parenchymal cells. The strategies and pipelines presented here for the liver are readily adaptable to other organs as well17,18,19.

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Protocol

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All mouse experiments used in this study have been approved by the Icahn School of Medicine at Mount Sinai Institutional Animal Care and Use Committee. The reagents and the equipment used are listed in the Table of Materials.

1. Generation of mice with tdTomato (TdT)-labelled HSCs

  1. Set up Tg(lox-stop-lox-TdTomato) x Tg(Lrat:Cre) breeders (Jackson Laboratories strain #00791420 and Mutant Mouse Resource & Research Centers (MMRRC) Strain #06959521-JAX, respectively).
    NOTE: Expect litter after 3 weeks. Use LratCre females for breeding, as some Cre expression has been detected in the male germline.
  2. Collect tails and genotype F1 litter to identify Tg(Lrat:Cre);Tg(LSL-TdT) mice that selectively express the TdTomato fluorophore in hepatic stellate cells.

2. In situ liver perfusion and fixation

  1. Pre-surgery set-up
    1. Prepare a ketamine/xylazine mixture for mice anesthesia by mixing 2 mL of ketamine, 1 mL of xylazine, and 7 mL of water.
      NOTE: Ketamine is a controlled substance and requires proper licensure and regulation for laboratory usage.
    2. Prepare the perfusion solution of 4% paraformaldehyde (PFA) in 1x Phosphate Buffered Saline (PBS).
    3. Set up the peristaltic pump by allowing 1x PBS to flow through the tube and set the flow rate at 2 mL/min.
      NOTE: Ensure that there is no air bubble in the tube.
  2. Weigh the mouse to determine the amount of the ketamine/xylazine mixture to inject.
  3. Inject the ketamine/xylazine mixture intraperitoneally into the mouse.
    NOTE: Inject 200 µL of the mixture for every 20 g of mouse weight.
  4. Place the mouse in a container with a lid and wait approximately 20 min for the mouse to be fully anaesthetized.
    NOTE: Pinch the mouse toe to check for the absence of reflex. Ensure that the mouse is still alive when perfusion initiates.
  5. Transfer the mouse from the glass container to the surgery platform in the supine position.
    CAUTION: PFA is toxic and volatile; therefore, perform the perfusion procedure in a chemical hood. Collect and dispose of the PFA flow-through properly according to institutional biosafety guidelines.
  6. Pinch the palms of the mouse foot with forceps to ensure the mouse does not have any reflex/spasm movements.
  7. Stab 4 palms with needles to spread the mouse's abdomen.
  8. Spray the abdomen with 70% ethanol to wet the fur.
  9. Use forceps to pick up the skin in the middle of the abdomen, align with the hind legs, and start cutting the skin (both layers) to reveal the abdomen.
  10. Use PBS-wet cotton swabs to move intestines and other organs to the side and lift the left lobe.
  11. Carefully cut the membrane attached between the left lobe and the caudate lobe.
  12. Use a suture to tie off at the top base of the left lobe.
  13. Cut the left lobe below the suture knot and set the left lobe aside for tissue sample collection and processing, depending on experimental needs (such as directly snap-freezing in liquid nitrogen, embedding in Optical Cutting Temperature (OCT) compound, or formalin fixation).
  14. Use PBS-wet cotton swabs to move the liver and reveal the portal vein.
  15. Carefully insert the catheter needle into the portal vein at about halfway along the length of the portal vein.
    NOTE: To assist with the catheter insertion, lift the mouse's body by placing a forceps under the body to ensure a flatter surface. Carefully align the catheter needle as parallel as possible with the portal vein before insertion.
  16. Remove the catheter needle.
    NOTE: Ensure a backflow of blood through the catheter to indicate successful insertion.
  17. Start the perfusion pump at a flow rate of 2 mL/min, connect the tubing with the catheter, and perfuse the liver with 20 mL of 1x PBS.
    NOTE: Ensure that the catheter is completely filled with blood, and the tubing is completely filled with the perfusion buffer, before connecting them to ensure absolutely no air bubbles get introduced into the system.
  18. Clamp the pulmonary artery above the liver with a hemostat to stop the 1x PBS from going through the heart.
  19. Cut the inferior vena cava to release the pressure buildup.
  20. Adjust the pump speed to 5 mL/min.
    NOTE: Allow approximately 15 min to finish the perfusion.
  21. Stop the pump to change the reservoir to 4% PFA.
    NOTE: Make sure not to introduce air bubbles into the tube by stopping the pump when there is still liquid in the reservoir.
  22. Perfuse the liver with 25 mL of 4% PFA.
  23. Once the perfusion is complete, remove the liver from the surrounding organs by cutting the connecting tissues.
  24. Place the liver in a 50 mL conical tube, filled with 25 mL of 4% PFA.
  25. Put the tube horizontally on a bench rocking platform overnight at 4 °C at 1.5 rpm, to ensure solution movement.
  26. Wash the liver 3 times in 1x PBS on a bench rocking platform for 30 min each at room temperature at 1.5 rpm.
  27. Store the liver in 20 mL 1x PBS with 0.02% NaN3 at 4 °C.
    NOTE: Store the liver long-term at this stage, up to at least one year.

3. iDISCO+-based tissue clearing

NOTE: The original description of the iDISCO+ protocol is accessible at 17

Day 1: Sample dehydration

  1. Cut the liver samples into pieces with sizes approximately 0.5-1 cm3.
    NOTE: Cut different samples into different shapes for easier detection during imaging steps.
  2. Prepare 20% (vol/vol), 40%, 60%, and 80% methanol dilutions in water in separate glass bottles.
  3. Place liver samples into glass vials with screw tops (e.g., 4-7 mL).
    NOTE: Place up to 6 liver sample pieces in one 7 mL glass vial.
  4. Dehydrate the sample using the methanol dilution in H2O series, 20% (vol/vol), 40%, 60%, and 80% for 1 h each at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  5. Wash the sample with another 100% methanol on a wheel rotator at 8 rpm, with no centrifugation occurring for 1 h at room temperature.
  6. Chill the sample at 4 °C for 30 min to 1 h.
  7. Prepare 66% (vol/vol) DCM dilutions in methanol in a glass bottle.
    NOTE: Prepare new dilutions every time. Use glass cylinders to prepare the solution, as DCM can dissolve plastic, such as polystyrene.
  8. Incubate samples in 66% DCM/methanol solution overnight on a wheel rotator at 8 rpm at room temperature, with no centrifugation occurring.
    NOTE: Beware of samples floating to the top once the solution is added.

Day 2: Sample bleaching

  1. Wash the sample twice with 100% methanol on a wheel rotator at 8 rpm, with no centrifugation occurring for 1 h each time at room temperature.
  2. Chill the sample at 4 °C for 30 min to 1 h.
  3. Bleach the sample in chilled 5% H2O2 in methanol (1 volume of 30% H2O2 to 5 volumes of methanol), on a wheel rotator at 8 rpm, with no centrifugation occurring, at 4 °C overnight.
    NOTE: Prepare a new solution every time.

Day 3-5: Sample permeabilization

  1. Rehydrate with methanol/H2O series: 80%, 60%, 40%, 20%, 1x PBS for 1 h at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  2. Wash the sample twice in PTx.2 (Table 1) for 1 h each at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  3. Incubate the samples in Permeabilization Solution (Table 1) at 37 °C for 2 days.

Day 5-17: Immunolabeling

  1. Block the samples in Blocking Solution (Table 1) for 2 days at 37 °C.
    NOTE: Prepare a fresh blocking solution every time due to the instability of donkey serum. Store leftover blocking solution at 4 °C for up to 1 month.
  2. Incubate the samples in primary antibodies diluted in PTwH (Table 1) with 5% DMSO and 3% Donkey Serum at 37 °C on a wheel rotator at 5 rpm, with no centrifugation occurring for 4 days.
    NOTE: Start with a 1:500 primary antibody dilution.
  3. Wash the samples in PTwH 4-5 times for 1 h each at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
    NOTE: Optionally, leave the samples to wash overnight.
  4. Incubate the samples in secondary antibodies and propidium iodide diluted in PTwH with 3% Donkey Serum at 37 °C on a wheel rotator at 5 rpm, with no centrifugation occurring for 4 days.
    NOTE: Start with a 1:500 secondary antibody dilution. Use a 1:250 propidium iodide dilution from a 5 mg/mL propidium iodide stock solution.
  5. Wash the samples in PTwH 4-5 times for 1 h each at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  6. Leave the sample washing in PTwH overnight, at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.

Day 18: Clearing

  1. Dehydrate the sample using the methanol dilution in H2O series, 20% (vol/vol), 40%, 60%, 80% for 1 h each at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  2. Wash the sample twice with 100% methanol on rotation for 1 h at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  3. Incubate the sample in 66% (vol/vol) DCM dilutions in methanol for 3 h at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  4. Wash the sample twice in 100% DCM for 15 min at room temperature on a wheel rotator at 8 rpm, with no centrifugation occurring.
  5. Incubate the sample in DBE overnight at room temperature before imaging.
  6. Store the samples long-term in DBE up to 1 year without impacts on the staining signal.

4. Confocal microscopy imaging

  1. Place the cleared sample in a 35 mm glass-bottom dish with a #1.5 glass bottom.
  2. Fill the dish with ethyl cinnamate to completely submerge the sample.
  3. Place the dish on the confocal and adjust the focus to identify the best plane that gives the brightest signal.
  4. Acquire a tile image on the brightest plane to capture a large area of the sample, using 1024 x 1024 format and speed 600.
    NOTE: Use a lower objective (10x or 20x) for the tile image to decrease imaging time and file size.
  5. Based on the tile image, select regions of interest to collect a higher resolution z-scan.
  6. Capture a z-scan through the cleared tissue, covering the whole depth of the tissue at the best resolution of the microscope, using 1024 x 1024 format and speed 600.
    NOTE: Use a 40x objective in oil immersion to give the best resolution for HSC.
  7. Save image files in LIF format and view files using Fiji or ImarisViewer 11.0.0.

5. Image analysis for 3D reconstruction and quantitative analysis of individual HSCs

  1. Convert image files to IMS format using the ImarisFileConverter 10.2.0.
  2. Use the Surface model on Imaris 10.2.0 to detect the object in the images.
  3. Use the Create wizard to perform analysis on a selected 3D region of interest, then apply it to the entire image.
    NOTE: Use this to speed up the analysis process.
  4. Select the source channel, absolute intensity, and uncheck the smooth button to define the intensity threshold for the whole image.
  5. Modify the threshold to capture all the objects in the image.
  6. Use the number of voxels to filter out debris objects.
  7. Finalize the wizard, press the statistic button to obtain all the statistical measurements.

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Results

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Successful perfusion and fixing of the mouse liver are represented by a uniform pale, beige-yellow color of the liver, reflecting efficient removal of blood and penetration of the fixative (Figure 1A,B). Improper perfusion results in dark red or brown regions of blood clots, which reduce the optical clarity and lead to a high background signal in these sample regions (Figure 1C).

During tissue clearing, the tissue sam...

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Discussion

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This protocol describes a reproducible and adaptable sample preparation pipeline leading to the three-dimensional visualization and quantitative analysis of HSC morphology in intact liver tissue. The integration of genetic fluorescent cell labeling, in situ liver perfusion, and a modified iDISCO+ clearing method enables imaging of large tissue volumes at single-cell resolution while preserving the spatial architecture of the liver microenvironment.

A critical step for the success of t...

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Disclosures

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The authors have nothing to disclose.

Acknowledgements

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This work is supported by 1R01 DK136016-01 and 5P30CA196521 (NCI center grant). Microscopy and/or image analysis were performed at the Microscopy and Advanced Bioimaging CoRE at the Icahn School of Medicine at Mount Sinai. This research was supported in part by the Tisch Cancer Institute at Mount Sinai, P30 CA196521 - Cancer Center Support Grant.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Anti-RFP Antibody,rabbit, pre-adsorbedRockland600-401-379
4-Chamber 35mm glass bottom dish with 20 mm microwell, #1.5 cover glassCellvisD35C4201.5N
AnaSed Injection (xylazine injection)Akorn, Inc.NDC 593990110-2020 mg/mL
Conical centrifuge tubes: 50ml Falcon352098
Cotton swabsMedlineMDS202000
Curved Locking HemostatsFisher Scientific16-100-117
Cy3-conjugated AffiniPure Donkey Anti-Rat IgG (H+L)Jackson ImmunoResearch712-165-153
Delicate Operating Scissors 4.75" Curved Sharp/BluntRoboz Surgical Instrument
728501
Dibenzyl Ether (DBE), 98%Sigma Aldrich108014
Dichloromethane (DCM),  ≥99.8%Sigma Aldrich270997
Dimethyl sulfoxide (DMSO), ≥99.9%Sigma AldrichD8418
Donkey anti Rabbit IgG (H+L) Highly Cross Adsorbed Secondary Antibody, Alexa Fluor 647ThermoFisher ScientificA31573
Donkey SerumEMD MilliporeS30
Ethyl cinnamate, 99%Sigma Aldrich112372
Extra Fine Micro Dissecting Scissors; 24 mm BladeFisher ScientificNC9232515
F4/80 Monoclonal Antibody ratThermoFisher Scientific14-4801-82
FijiNIH
ForcepsFisher ScientificNC9214139
GlycineBio-Rad1610718
Heparin Sodium SaltChem-Impex29984
Hybaid Hybridization OvenTriad ScientificMaxi 14
Hydrogen peroxide, 30 %Sigma Aldrich31642
Imaris 10.2.0 softwareOxford Instruments
ImarisFileConverter 10.2.0 softwareOxford Instruments
ImarisViewer 11.0.0Oxford Instruments
IV CatheterExel International Inc.EXE 26751
Ketamine, Hydrochloride injectionDechraB8U4100 mg/mL
LAS X 4.4.0.24861 softwareLeica
Leica Stellaris 8 Confocal MicroscopeLeica
Methanol, HPLC gradeFisherA452-2
Multi-purpose rotatorScientific Industries, Inc.Model 151
Needles (27G x 1/2'')BD305109
Paraformadehylde, powder, 95%Sigma Aldrich158127
Peristaltic pumpGilsonModel Minipuls 3
Peristaltic pump tube, MasterFlexThermo Scientific96400-163.1mm ID x 7.6m L/S Peroxide-Cured Silicone Precision Pump Tubing, Size L/S 16, 1/8in Hose Barb
Phosphate Buffered Saline (PBS)CytivaSH30256.01Sterile, pH 7.0 - 7.2, without calcium, magnesium
Propidium iodide, ≥94.0% (HPLC)Sigma AldrichP4170
Razor BladesAmerican Line66-0362
Rocking platformVWRModel 200
Sodium Azide, ≥99.5%Sigma AldrichS2002
SutureEthiconJ109T
Syringe (1 mL)BD309628
TritonX-100SigmaT9284
Tween-20Thermo ScientificJ20605-AP
Vials, screw top with solid cap with aluminum liner, preassembledSigma Aldrich27470-Uvolume 7 mL, amber glass vial, O.D. × H 17 mm × 60 mm, thread for 15-425

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Tags

Hepatic Stellate CellsTissue ClearingConfocal MicroscopyThree Dimensional ReconstructionLiver Tissue ImagingFibrogenic MyofibroblastsFluorescent LabelingMouse Liver Models

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