Name: Reusable Hepatic Imaging Tool: A Reusable 3D-Printed Tool for Hepatic Intravital Microscopy (Video) | JoVE Journal
Uploaded: 2026-03-17T13:25:00.000+00:00
Duration: 11 min 29 s
Description: Learn how to fabricate and use the Reusable Hepatic Imaging Tool for high-resolution intravital microscopy of the liver in live mice. This 3D-printed device improves tissue stability and reduces motion artifacts during extended imaging sessions.

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5 minutes

This work was supported by the Intramural Research Program at the US National Institutes of Health, the National Cancer Institute, and the NIH library. H. Sanchez Cornejo thanks the Peruvian Agency CONCYTEC for financial support through grant No.PE501087367-2024-PROCIENCIA. We want to thank the members of the Porat-Shliom lab and collaborators at the University of Cambridge Quantum Physics group for critical reading, the NIH library staff, and Geneti Gaga for technical assistance. The 3D-rendered image in Figure 2A was created using Biorender.

All the animal studies were carried out at the animal facilities of the National Institutes of Health in compliance with guidelines and protocols authorized by the Ethics Committee for Animal Experiments (protocol No. LCMB-037). All personnel are required to be properly trained before proceeding with the following protocols. Animal handling was performed in compliance with institutional guidelines.
The RHIT protocol outlines the preparation of Reusable Hepatic Imaging Tools (RHITs) for use in intravital microscopy (IVM) experiments. The experimental procedures are performed in a sterilized microscope room, or hood, and involve a series of steps: first, the printing of RHIT, anesthesia, and fluorescent labeling of the mouse, then the setup of the microscope, next the surgical procedure, and finally the application of RHIT-assisted IVM. Figure 1 and Figure 2 illustrate the required components for imaging live specimens using a confocal microscope, which include a metal insert5,6,7 (microscope adaptor for imaging live specimens) for placement of RHIT, as well as RHITs for use in advanced or controlled steatosis. For testing the system, male C57BL/6J Mus musculus aged 6-10 weeks were obtained and fed a Standard or Western diet for 8-10 weeks prior to the experiment.
1. Designing RHIT with modifications
<ol>
	<li>To recreate RHIT, use 3D product design software (see the Table of Materials). Open the design software and create a new standard part. Save the file with an appropriate description.</li>
	<li>Create the sketch of RHIT on the front plane using the Create 2D Sketch feature.
	<ol>
		<li>Click on the rectangle icon and place its bottom-left corner at the origin of the sketch plane.</li>
		<li>Draw a second rectangle inside the first one to represent the imaging window of RHIT.</li>
		<li>Add dimensions to both rectangles using the Dimension feature, ensuring all lengths and widths are referenced to the origin and relative to each other.</li>
		<li>Adjust the RHIT dimensions according to the mouse size.</li>
		<li>Add thickness to RHIT by selecting the outer rectangle only and clicking extrude. When a popup appears, select the sketch using the Extrude tool to leave an opening in the center.</li>
		<li>Select the bottom face of the extruded RHIT and start a new 2D sketch using the Start 2D Sketch feature.</li>
	</ol>
	</li>
	<li>Draw a rectangle to define the pocket for the heat pad. Apply the acquired dimensions using the Dimension feature.
	<ol>
		<li>Exit the 2D sketch and perform a material removal using the Cut Extrude tool to create a deep cavity for the heat pad.</li>
	</ol>
	</li>
	<li>Start a new sketch for the temperature probe on the same face as the cavity. Click on start a 2D sketch and select the circle icon. Place the circle on the plane surface and dimension it accordingly. Exit the sketch mode.
	<ol>
		<li>Click on extrude and select the sketch of the circle. In extrusion mode, click on cut extrude, and wait for a popup to appear with a through all option to remove material. Click OK. Exit extrusion mode.</li>
		<li>Add fillets on the edges of the window by clicking on the fillet icon and selecting all eight sharp edges. Add fillet size and exit fillet mode. 
		​NOTE: If additional motion absorption is desired, users may increase the internal infill density during 3D-printing preparation.</li>
	</ol>
	</li>
	<li>Prepare the model for 3D printing by navigating to the environment tab and selecting 3D Print tab.
	<ol>
		<li>Use the Set Orientation function to ensure the model is lying flat on the print bed and not oriented vertically.</li>
		<li>Click on Print Options to configure unit settings and adjust the scale if needed.</li>
		<li>Save the finalized model in. STL format using the STL export function.</li>
		<li>Exit the environment tab and click on save to save RHIT as a solid part.</li>
	</ol>
	</li>
	<li>Close the design program. 
	NOTE: Ensure the printer is compatible with the .STL file format or convert to the appropriate format as required by the 3D printer software.</li>
</ol>
2. Uploading and printing RHIT from STL files
<ol>
	<li>Import the .stl file from Figure 1A or Supplemental File 1, into 3D computer-aided design (CAD) software by clicking Import and selecting the .stl file. Click on the Prepare button to slice the model into thin layers for 3D printing.</li>
	<li>Select the appropriate 3D commercial printer and click on Print. Then, monitor the print progress, and once the print is complete, remove and clean the 3D printed object as needed.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Mouse Size</td>
			<td>Mouse Body Weight (g)</td>
			<td>RHIT Thickness (mm)</td>
			<td>Imaging Window Size L X W (mm)</td>
		</tr>
		<tr>
			<td>Small (S)</td>
			<td>17-20</td>
			<td>1.5</td>
			<td>17 x 17</td>
		</tr>
		<tr>
			<td>Medium (M)</td>
			<td>22-25</td>
			<td>2.5</td>
			<td>21 x 21</td>
		</tr>
		<tr>
			<td>Large (L)</td>
			<td>35-40</td>
			<td>3</td>
			<td>26 x 26</td>
		</tr>
	</tbody>
</table>
Table 1: Scaling mouse size to RHIT. The mouse weight is scaled to the thickness of RHIT, where a weight greater than 30 g (arrowhead) corresponds to the largest RHIT (3.0 mm). The lightest mouse corresponds to the thinnest RHIT, and the obese mice are represented by the thickest RHIT. These dimensions can be adjusted in CAD software. RHIT length and imaging aperture can vary by the organ location of the specimen being used; constant length and imaging aperture for all three RHITS were suitable for this study. Abbreviations: RHIT = Reusable Hepatic Imaging Tool; CAD = computer-aided design.
3. Microscope preparation and utilizing RHIT
<img alt="Microscopy setup in metabolic research; obese vs lean mice, tissue analysis, fluorescence imaging." class="xfigimg" src="/files/ftp_upload/68337/68337fig2.jpg" data-altupdatedat="1776139283000" data-alt="Figure 2"> 
Figure 2: IVM of liver using the RHIT technique. (A) Schematic of the inverted confocal microscope setup with an anesthetized mouse. (B) A metal microscope adaptor (insert) was used to position the mouse on the microscope stage for live imaging. (C) The 3D-printed RHIT was used with a gauze strip to easily manipulate liver parenchyma. (D) Image of the pre-exteriorized liver before RHIT positioning. The nose cone was used for anesthesia prior to and during surgery. (E) Image of the exteriorized liver prior to RHIT positioning during an acute procedure. Shaving was not required. (F) Image of the inverted mouse in the final face-down position on the RHIT for imaging. (G) RHIT securely attached to the microscope stage. (H) A mouse positioned on RHIT and metal adaptor. (I) Lean (blue arrow) and obese (green arrow) mice shown with respective scaled RHIT inserts in J-L depict gross anatomical images of the obese and lean liver (respectively). The high-resolution IVM images M,N of liver from obese and lean mice labeled with BODIPY (green, depicted by a pink arrow) to label lipid droplets and Hoechst 33342 (white, depicted by a cyan arrow) to label nuclei, respectively. Abbreviations: RHIT = Reusable Hepatic Imaging Tool; IVM = intravital microscopy. <a href="https://www.jove.com/files/ftp_upload/68337/68337fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Prepare a conventional inverted confocal microscope for liver IVM as shown in Figure 2.
	<ol>
		<li>Insert a stage adaptor to accommodate the sample. To follow this protocol, use a custom-made metal insert with a 35 mm circular opening and a 40 mm cover glass.</li>
		<li>Thoroughly clean both sides of the cover glass before and after each imaging using 70% ethanol as necessary.</li>
	</ol>
	</li>
	<li>Secure the appropriate RHIT based on mouse size (Table 1) with tape (optional) by aligning the RHIT aperture with the objective lens and parallel over the imaging coverslip (Figure 2G) on the metal insert with tape over the cover glass.</li>
	<li>Choose an objective for imaging based on the on-area size, spatial resolution, and the imaging time, such as a 25x/0.95 water objective. 
	NOTE: While higher magnification improves resolution, it also increases the likelihood of motion artifacts.
	<ol>
		<li>Level the objective close to the cover glass.</li>
	</ol>
	</li>
	<li>Secure a strip of gauze (approximately 1 x 8 cm, Figure 2C,D) across the RHIT window with tape to help position the tissue and separate it from other abdominal organs (Figure 2C).</li>
	<li>Prewarm RHIT using a heat pad inserted into the slot and secure the heat probe adjacent to the RHIT (Figure 2G,H) to maintain the body temperature of the mouse during imaging.</li>
</ol>
4. Fluorescent labeling and mouse anesthesia
<ol>
	<li>Reduce the mouse's stress by sanitizing the anesthetic induction chamber to get rid of any last traces of animal stench.</li>
	<li>Turn on the oxygen condenser, an environmental protection system (EPS), and the isoflurane vaporizer.</li>
	<li>Place the mouse in the induction chamber and anesthetize using 2-3% isoflurane for approximately 3-5 min, or until the animal exhibits stable respiration and loss of righting and pedal reflexes.
	<ol>
		<li>Once anesthetized, transfer the mouse to the heating pad and maintain anesthesia using 1-2% isoflurane delivered through the nose cone (Figure 2D), adjusting as needed to ensure a stable surgical plane of anesthesia throughout the procedure.</li>
		<li>Assess the depth of anesthesia by doing a toe pinch test to observe any voluntary movement.</li>
	</ol>
	</li>
	<li>To label lipid-rich structures, inject 200 nmol of BODIPY 665/676 diluted in PBS (total volume 200 µL) via the tail vein using a 31 G insulin syringe.
	<ol>
		<li>For optimal signal, administer BODIPY once daily for 2 consecutive days prior to imaging. Allow at least 3-4 h after the second injection to ensure systemic clearance of background and stabilization of tissue signal.</li>
	</ol>
	</li>
	<li>To provide nuclear contrast, inject Hoechst 33342 (2 µg/g body weight) diluted in 1x heparin solution (prepared by mixing 120 µL of heparin with 180 µL of saline to yield 400 U/mL) via the retro-orbital sinus using a 31 G insulin syringe. 
	NOTE: Hoechst is administered separately to avoid precipitation when mixed with other dyes.</li>
	<li>To visualize the vasculature, inject 18 µg/g body weight of fluorescent dextran (70 kDa) diluted in the same 1x heparin solution described in Step 4.5, via the contralateral retro-orbital sinus using a 31 G insulin syringe (Table 2). 
	​NOTE: Heparin is included to reduce clot formation and promote consistent blood flow during imaging.</li>
	<li>Wait 10-15 min after Hoechst and dextran injections before starting the surgery and imaging.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Fluorescent Reporter</td>
			<td>Purpose</td>
			<td>Dose per weight (or Concentration)</td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Stain nuclei</td>
			<td>2 µg/g: supplement injectant volume to 50 µl with heparin solution</td>
		</tr>
		<tr>
			<td>Bodiby 665/676</td>
			<td>Stain lipid droplets</td>
			<td>.8 µg/g: supplement injectant volume to 200 µl in saline solution</td>
		</tr>
		<tr>
			<td>2M tetramethyl rhodamine-dextran (Dextran)</td>
			<td>Stain vasculature</td>
			<td>18 µg/g: supplement injectant volume to 50 µl with heparin solution</td>
		</tr>
	</tbody>
</table>
Table 2: Fluorescent reporter options. This table provides a detailed description of the methods used to label mouse tissues for in vivo imaging. The heparin solution is prepared first and then used to make the DAPI-heparin and dextran-heparin solutions7. Hoechst 33342 is injected retro-orbitally into one eye, while BODIPY is injected directly into the tail vein 2-3 days prior, with a higher concentration for obese mice. Dextran is injected retro-orbitally into the other eye. It is important to perform retro-orbital injections separately in each eye. Abbreviation: DAPI = 4',6-diamidino-2-phenylindole.
5. Mouse surgical procedure
<ol>
	<li>Shave the abdominal coat using a hair clipper and/or hair removal cream. Remove hair residue and use 70% ethanol to clean the exposed skin. 
	NOTE: Depending on institutional IACUC requirements, an alternating scrub of betadine and ethanol may be used to ensure appropriate surgical site preparation.</li>
	<li>Reassess the depth of anesthesia prior to surgery.</li>
	<li>Expose the liver by making a horizontal crescent-shaped 2-3 cm incision under the ribcage with surgical scissors from the xiphoid, ending at the left lateral side of the upper abdomen.</li>
	<li>After the initial incision, carefully make a smaller secondary incision in the muscle layer to expose the left lobe of the liver, cauterize (optional) as necessary to avoid bleeding (Figure 2E).
	<ol>
		<li>Prevent heat damage to the liver whilst using the cauterizer by placing a saline-soaked gauze between the liver and muscle layer to absorb the emanating heat of the muscle layer.</li>
		<li>Avoid any excessive bleeding, surgical or mechanical trauma, or external compression.</li>
	</ol>
	</li>
	<li>Once the liver is exposed, transfer the mouse to the microscope stage.</li>
</ol>
6. Mouse placement on the microscope stage
<ol>
	<li>Position the anesthetized mouse's abdomen facing down on top of the RHIT platform of the microscope.
	<ol>
		<li>Adjust the nose cone so that the mouse remains under anesthesia.</li>
		<li>Employ a saline-moistened cotton-tipped applicator to delicately maneuver the left lateral lobe of the liver against the coverslip while applying gentle external pressure to the thoracic cavity. 
		NOTE: Take advantage of gravity to carefully maneuver the liver out and onto the cover glass.</li>
		<li>Use a gauze strip to gently separate the liver from the mouse's body ( Figure 2E-H).</li>
	</ol>
	</li>
	<li>Check the nose cone again to ensure the mouse remains under anesthesia during imaging.</li>
	<li>Prevent tissue dehydration by applying a water-based gel around the edges of the exposed cavity and avoiding direct contact with the heat pad. 
	​NOTE: Maintaining temperature and humidity can also be aided by a microscope with an enclosed humidity chamber5,6,7.</li>
	<li>Apply ophthalmic ointment to the eyes to prevent corneal drying.</li>
	<li>Set up time-lapse IVM to begin imaging7,52.
	<ol>
		<li>Open laser shutters, evaluate fluorescent labeling, and identify a stable region of interest with a notable blood flow. Select appropriate image acquisition parameters, including image format, acquisition speed, imaging intervals, total acquisition duration, and line/frame averaging (512 × 512 pixels; 1-5 frames s⁻¹ per region; imaging intervals of 5-30 s; total acquisition time of 5-20 min; line/frame averaging of 2-8, depending on experimental requirements). Optimize fluorescence flash exposure to accommodate the fluorophores used.</li>
		<li>Define upper and lower boundaries for Z-stacks by selecting Z-Stack in the acquisition panel, adjusting the top and bottom focal planes using the Z-position controls, and confirming the range with Set First/Set Last; then initiate the time-lapse recording by clicking Start Experiment (or Start Acquisition). At the conclusion of imaging, euthanize the animal in accordance with institutional animal care and use committee-approved protocols.</li>
	</ol>
	</li>
</ol>
<img alt="Fluorescent labeling microscopy images with Bodipy, Hoechst, Dextran, and SSIM graph analysis." class="xfigimg" src="/files/ftp_upload/68337/68337fig3.jpg" data-altupdatedat="1776139283000" data-alt="Figure 3"> 
Figure 3: Measuring the improved hepatic IVM stability of RHIT with SSIM. Hepatic IVM of the lower left liver lobe under three stabilization conditions: (A) no insert (NONE), (B) cardboard insert, and (C) RHIT. Tissues are labeled with BODIPY (magenta, lipids), Hoechst 33342 (green, nuclei), and Dextran (cyan, vasculature); panels A and B show visible distortion and motion artifacts, whereas C demonstrates enhanced clarity with RHIT. (D) Structural similarity index (SSIM) was computed to quantify image stability. Bars show mean ± SEM, and each colored dot represents an individual biological replicate: RHIT (n = 14), Cardboard (n = 10), No Insert (n = 8). These groups are subsets drawn from the overall experimental cohort of approximately 50-70 mice. Only sessions meeting specific imaging criteria (e.g., consistent frame alignment and image quality) were analyzed via SSIM. RHIT consistently produced higher SSIM values (mean 0.511 ± SEM), indicating improved stability and resolution across imaging conditions. Scale bars = 15 µm. Representative images are shown; SSIM reflects serial IVM sessions without observable tissue damage or disruption. Abbreviations: RHIT = Reusable Hepatic Imaging Tool; IVM = intravital microscopy; SSIM = structural similarity index; CARD = cardboard insert. <a href="https://www.jove.com/files/ftp_upload/68337/68337fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors have no conflicts of interest to declare.

Liver intravital microscopy has been extensively studied, however, previous methods often require highly invasive and unstandardized surgical procedures for stabilized terminal imaging5,6,7,9,31,32. Motion is arguably the most challenging barrier preventing reproducible imaging in hepatic IVM. The most effective solution for ...

Intravital microscopy is a powerful tool to visualize biological processes in vivo dynamically1,2,3 in contrast to the traditional immunofluorescence histology with fixed tissue or live cell imaging and flow cytometry analysis of isolated cells. For liver inspection, IVM has been utilized to examine bile secretion and flow4,5, glucose mobilization6,7, nonparenchymal cells8,9,10 and liver diseases11,12. Moreover, with regards to lipid accumulation investigation, IVM allows real-time information on the dynamics of lipid droplet formation, growth, and degradation in live tissues such as the visualization and quantification of lipid droplets at high spatial resolution, and for the direct correlation of lipid droplet accumulation with other cellular and physiological processes13. Despite these advantages, IVM is still technically challenging: it requires specialized equipment and expertise14, can cause tissue damage, alter cellular behavior, and demand invasive procedures. Traditional in vivo studies relying on behavioral observation, physiological monitoring, and genetic analysis provide valuable information but lack the spatial and temporal resolution offered by intravital microscopy.
The first major challenge with IVM is controlling the instability of the artifacts caused by heartbeat and respiration of the inspected animal, which can compromise the image quality, resulting in obscured or distorted images of the biological processes being observed15. Attempts to correct this with image processing often result in losing important spatial and temporal information or demanding advanced surgical equipment and maintenance16,17. Secondly, mouse models for liver disease studies, such as MAFLD and T2D, are often obese and, thus, harder to stabilize due to their increased size and tissue lipid content16,18. In obese NAFLD and T2D mouse models, increased body mass poses mechanical challenges for intravital imaging by exceeding the load capacity of standard microscope stages and complicating animal stabilization19. This added weight increases the risk of motion artifacts and tissue compression, both of which impair imaging precision20.
Simultaneously, elevated hepatic lipid content increases tissue opacity and light scattering, which degrades image resolution and contrast. These effects are often compounded by metabolic comorbidities such as altered perfusion and inflammation, further reducing dye distribution and signal clarity21. Finally, it can be difficult and impair repeatability to maintain a steady body temperature throughout imaging to sustain tissue physiology and biological activities. In addition, stable positioning of the liver on the microscope stage during IVM is crucial to prevent mechanical stress or tissue damage7. Exposing the liver tissue is a delicate procedure that requires technical precision to prevent severe bleeding and liver parenchyma damage7. In order to reduce motion artifacts, the mouse's ribcage was previously lifted from the microscope stage using cardboard, allowing gravity to drag the liver away from the rest of the body22. Although this approach was shown to be successful in producing quantifiable data, the cardboard material is unsustainable as it loses shape and is difficult to clean. Additionally, it is hard to change the dimensions of the cardboard insert to accommodate obese mouse models.
In recent years, several advanced hepatic intravital microscopy techniques have emerged, including scanning light field microscopy and computationally enhanced optical systems, which have demonstrated improvements in spatiotemporal resolution23, reduced phototoxicity24, greater imaging depth25,26, and motion artifact suppression27. These developments, as demonstrated in recent studies28,29,30, represent important technological progress in the field. This paper presents the Reusable Hepatic Imaging Tool (RHIT), an innovative tool for IVM of the murine liver. RHIT contributes to this evolving landscape by offering a scalable and reusable tool that enhances mechanical stability and tissue accessibility across diverse animal models, including those with metabolic disease. Its design addresses the persistent challenges of motion, variability, and setup complexity by integrating anatomical fit, staging reproducibility, and ease of use. RHIT's compatibility with both conventional and advanced imaging modalities expands the utility of these emerging techniques, promoting reliable and physiologically stable intravital liver imaging.
While various approaches to stabilize the tissue during IVM have been proposed, such as a surgical window9,31,32,33,34,35,36 and custom assembled acrylic stage6,37, these methods also introduce numerous limitations and complexities with regard to scalability, reproducibility, and surgical skills. Comparatively, RHIT is functionally distinct in its purpose and design. It does not rely on direct compression or gel-based embedding38. Instead, RHIT serves as a scalable, reusable, anatomically contoured tool that supports liver isolation via gravitational positioning, maintaining physiological orientation without restricting or distorting the tissue. Critically, RHIT improves data reproducibility, imaging fidelity, and procedural consistency across a wide range of mouse sizes. It eliminates the need for gels, adhesives, or complex surgical interfaces, while simplifying tissue alignment and temperature control. Moreover, RHIT is a specialized tool that bridges the interface between the animal and the microscope, engineered to assist with both physiological support and imaging precision.
RHIT directly addresses the mechanical and optical challenges of metabolically compromised models through its scalable geometry, which accommodates larger animal sizes while evenly distributing weight to minimize platform instability. The recessed liver aperture stabilizes the tissue using gravity without compressive force, enabling consistent access, and reducing motion during imaging. Its reusable, chemically inert design ensures compatibility with biological tissues while supporting hygienic reuse. By enabling consistent positioning, minimizing motion artifacts, and maintaining tissue integrity, RHIT provides a foundational advancement in intravital microscopy workflows. Its adaptability and open-source availability further promote broad utility across laboratories and animal models, setting a new standard for imaging tools that support both the specimen and the imaging system.
Furthermore, commercially available heated microscope stages may require advanced microscope setups and handling39. They are not scalable to the specimen and are generally more expensive, making them less accessible for many researchers. Thus, a reusable scalable insert is important to standardize IVM for different animal models. 3D printed technology, also referred to as additive manufacturing, has become increasingly popular in scientific laboratories40, transforming biochemistry, physics, and biology lab protocols. For IVM, 3D technology has been used to manufacture lightweight imaging optical windows41,42, improve tissue accessibility43, and facilitate tissue repair and reconstruction44. Thus, a customized tool for tissue stabilization, which, to our knowledge, has not been described, poses an excellent opportunity to employ 3D printing to overcome some of the challenges of hepatic IVM.
RHIT is created utilizing a one-step 3D printing method and substantially improves tissue stabilization for an extended period. RHIT is highly accessible and suitable for acute IVM experiments that require minimal surgical expertise. In addition, RHIT is easily scalable to animal size and has a low cost of production. Finally, we demonstrate that RHIT enables stable time-lapse fluorescence imaging and improves tissue stability compared to the previously used cardboard insert. RHIT simplifies IVM of the hepatic lipid homeostasis without direct contact or measurable impact on physiological conditions. Consequently, RHIT presents a competitive edge as an accelerated, reusable, and adaptable contribution to IVM34,36,45.
<img alt="Mouse liver aperture design; engineering schematic and 3D model with probe insert." class="xfigimg" src="/files/ftp_upload/68337/68337fig1.jpg" data-altupdatedat="1776139283000" data-alt="Figure 1"> 
Figure 1: RHIT design. (A) Design and dimensions of a 3.0 mm RHIT scaled for an obese mouse. The enface view (left) shows the dimensions, positioning, and size of the tissue aperture, as well as the orientation of the mouse during imaging. The side view (right) includes dimensions and indicates the position of the heat pad and temperature probe insert. Scaling parameters for different mouse sizes are provided in Table 1. (B) 3D enface view of RHIT generated from the open-source STL file, showing where the rectal probe wire is inserted through the custom port (labeled rectal probe insert dimensions shown upper right). (C) Technical assembly of RHIT showing integration of the heating device and the custom structural inserts for various RHIT sizes, the wire is fitted into the rectal probe insert for temperature probe access. Abbreviation: RHIT = Reusable Hepatic Imaging Tool. <a href="https://www.jove.com/files/ftp_upload/68337/68337fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
RHIT was originally designed as a single-part assembly using ABS (Acrylonitrile Butadiene Styrene), a widely used thermoplastic polymer in fused deposition modeling 3D printing45. ABS offers several advantages for this application: it is lightweight, heat-resistant (melting point ~200 °C), and has good impact resistance46,47,48. RHIT's design takes advantage of the tunable density of ABS, allowing it to absorb physiological motion such as heartbeat and respiration. Increased infill density enhances motion dampening, resulting in greater imaging stability and improved image quality.
RHIT is designed to absorb motion through adjustments to the density of a chemically inert material such as ABS. In this context, density refers specifically to the internal infill percentage used during 3D printing, which affects the rigidity and ability of the platform to dampen vibration49,50,51. Increasing the density of ABS has been observed to reduce the transmission of movement caused by heartbeat and respiration, resulting in a stable platform for imaging and improved image quality. We tested RHIT constructs with varying internal fill densities (20-100%) and observed qualitatively that higher-density configurations led to visibly reduced motion artifacts during liver intravital imaging in mice. Formal quantification was not performed, as the high-density configuration proved sufficiently sturdy and effective across all RHIT sizes tested, and consistently provided reliable performance under routine imaging conditions. For most users, this default density will be suitable without further adjustment. However, for experiments requiring specific mechanical characteristics, the infill density and design parameters can be modified, and full 3D printing instructions have been provided to support such customization.
Alternative materials can be used for printing; however, ABS is the preferred material for RHIT fabrication as it offers several key advantages. It is chemically inert, minimizing the risk of interaction with tissue or bodily fluids, and insoluble in water, enabling efficient cleaning, storage, and reuse. These properties make ABS a reliable and durable choice for experimental use. When access to a fused deposition modeling printer is limited, other 3D printing techniques and materials can be used. Stereolithography, selective laser sintering, and multi-jet fusion are suitable options with materials such as nylon, photopolymer resins, or composite powders. Nylon provides durability and flexibility, whereas stereolithography resins offer high resolution for applications requiring fine structural detail. The RHIT design can be readily adapted to the tolerances and constraints of different materials, making it compatible with a range of printing platforms. Users should select materials based on mechanical stability, biocompatibility, compatibility with imaging workflows, and available printing resources.

<table><tbody><tr><td>&amp;frac12;-inch masking tape</td><td>3M General Purpose Masking Tape:</td><td>b40069428</td><td></td></tr><tr><td>40-mm coverslips</td><td>Bioptech</td><td>40-1313-03191</td><td></td></tr><tr><td>70% (v/v) ethanol</td><td>Thermo Fisher Scientific</td><td>BP2818-4</td><td>70% ethanol by volume and 30% water by volume</td></tr><tr><td>Anesthesia machine: AMD-3 Plus</td><td>Somni</td><td>AMD-17106</td><td></td></tr><tr><td>Aquasonic 100 ultrasound transmission gel</td><td>Parker Laboratories</td><td>&amp;nbsp;01 08</td><td></td></tr><tr><td>BODIPY 665/67</td><td>Thermo Fisher Scientific (Invitrogen)</td><td>B3932</td><td>See Table 2</td></tr><tr><td>CAD software (Autodesk Inventor or equivalent)&amp;nbsp;</td><td>Autodesk (representative CAD software)</td><td>n/a</td><td>Software</td></tr><tr><td>Cautery System</td><td>Gemini</td><td>GEM 5917</td><td></td></tr><tr><td>Clidox disinfectant</td><td>Pharmacal Research Laboratories Inc.</td><td>VWR, MSPP-96118F</td><td></td></tr><tr><td>Confocal microscope (Leica SP8 or equivalent)</td><td>Leica</td><td>SP8</td><td></td></tr><tr><td>Cotton-tip applicators</td><td>Puritan Medical Products</td><td>25-806 1WC</td><td></td></tr><tr><td>Custom-made aluminum stage insert (3 mm thick, 35-mm depression)</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>FORANE (isoflurane, USP) liquid for inhalation, 250 mL</td><td>Baxter</td><td>10019-360-60</td><td></td></tr><tr><td>Hair clipper (optional)</td><td>Onyx Fine Instruments</td><td>ASWRHC</td><td></td></tr><tr><td>Heat pad with rectal probe</td><td>ALA Scientific Instruments</td><td>HEATINGPAD-1/2</td><td></td></tr><tr><td>Heparin sodium, 1000 USP U/mL</td><td>Hikma</td><td>0641-0391-12</td><td></td></tr><tr><td>Hoechst 33342</td><td>Thermo Fisher Scientific (Invitrogen)</td><td>62249</td><td>See Table 2</td></tr><tr><td>Mouse heat pad</td><td>ALA Scientific Instruments</td><td>HEATINGPAD-1/2</td><td></td></tr><tr><td>Mus musculus</td><td>The Jackson Laboratory</td><td>C57BL/6</td><td>Laboratory mouse</td></tr><tr><td>Normal saline (0.9% sodium chloride)</td><td>Thermo Fisher Scientific</td><td>7210-16</td><td></td></tr><tr><td>RHIT (Reuseable Hepatic Imaging Tool) applicable size</td><td>Provided</td><td>N/A</td><td>See Table 1</td></tr><tr><td>Surgical forceps, curved tip</td><td>Fine Science Tools</td><td>11251-23</td><td></td></tr><tr><td>Surgical gauze</td><td>Mckesson</td><td>6939</td><td></td></tr><tr><td>Surgical scissors</td><td>Fine Science Tools</td><td>14001-14</td><td></td></tr><tr><td>Tetramethylrhodamine Dextran</td><td>Thermo Fisher Scientific (Invitrogen)</td><td>D7139</td><td>See Table 2</td></tr><tr><td>Ultra Fine U-100 Insulin Syringes, 31 G</td><td>Betty Mills</td><td>BD326730</td><td></td></tr></tbody></table>

reusable hepatic imaging tool: a reusable 3d-printed tool for hepatic intravital microscopy

Reusable Hepatic Imaging Tool: A Reusable 3D-Printed Tool for Hepatic Intravital Microscopy

Hepatic function and dysfunction can be assessed by directly visualizing the dynamic processes of the liver using confocal-based intravital microscopy (IVM). In particular, excellent spatial and temporal precision can be achieved in the extraction of both qualitative and quantitative information. On the microscope stage, maintaining physiological tissue conditions while achieving stability and repeatability remains challenging. Increased weight and hepatic lipid content have a detrimental effect on this process in animal models of Type II diabetes (T2D) and non-alcoholic fatty liver disease (MAFLD). To overcome these constraints, we have created RHIT, a revolutionary imaging instrument design that improves tissue access and stability. RHIT is a reusable, scalable hepatic imaging device manufactured using 3D printing in an efficient one-step method. Its dimensions may be changed to fit mice of different sizes. Briefly, RHIT is a 3D-printed apparatus for positioning and stabilizing laboratory animals during intravital liver microscopy, used after a brief surgical preparation to minimize handling, stress, and variability. Its reusable, standardized design is easily produced and customized, supporting a range of sizes and potential experimental designs. Initial results demonstrate a notable improvement in the stability and reliability of hepatic intravital imaging, enabling the detailed visualization of subcellular structures, such as lipid droplets. This improved precision facilitates metabolic and physiological studies with improved clarity, reproducibility, and potential for longitudinal designs. The protocol offers a standardized, accessible approach that addresses long-standing technical challenges in hepatic IVM.

RHIT was designed using a CAD application (Figure 1). RHIT dimensions are scaled to accommodate small, medium, and heavy-sized mice (Table 1). RHIT design consists of a 107.50 mm long ABS flat material with an aperture positioned to accommodate liver imaging. IVM was performed under optimal (RHIT) and suboptimal (no insert or cardboard) conditions (see Figure 3A-C). This illustrates the range of possible outcomes and the extent ...

This protocol introduces a cutting-edge, Reusable Hepatic Imaging Tool (RHIT) for intravital microscopy (IVM) of live mice, which enhances tissue stability for rapid and acute experiments. The RHIT&#39;s 3D-printed design offers cost-effective production, scalability, and versatility, enabling prolonged observation of metabolic processes in mice of various sizes.

Hepatic function and dysfunction can be assessed by directly visualizing the dynamic processes of the liver using confocal-based intravital microscopy ...

The overall goal of this procedure is to showcase the Reusable Hepatic Imaging Tool, or RHIT, as a breakthrough tool for capturing high-quality, dynamic biological data, especially in liver research. Traditional scientific methods for studying mice in controlled experiments have long relied on behavioral observation, physiological monitoring, and genetic analysis. The complexity of these studies frequently results in inconsistent or limited data outputs.Many experiments rely on data that is either incomplete or difficult to interpret, such as brief flashes of low-resolution images that quickly fade, making it hard to capture long-term trends or subtle changes. Introducing RHIT, enabling dynamic, high-resolution capture for liver research and beyond. The technique we demonstrate enables the fabrication of custom imaging tools using accessible materials and widely available 3D printing technology.Designed for use in academic and resource-limited environments, this approach allows for the rapid development and deployment of devices to support high-resolution in vivo imaging. Our Reusable Hepatic Imaging Tool, or RHIT, was developed to improve tissue stability during intravital microscopy of the liver. RHIT's modular design accommodates mice of various sizes and supports extended imaging of dynamic metabolic processes with minimal physiological disruptions.RHIT is fabricated in a single step using acrylonitrile, butadiene, styrene, ABS, a thermoplastic selected for its light weight, thermostability, and mechanical durability. By adjusting material density, RHIT effectively absorbs motion caused by respiration and heartbeat, thereby reducing image artifacts and enhancing clarity. ABS's chemical inertness and water insolubility ensure availability with biological tissue and enable efficient cleaning and reuse.To recreate RHIT, use a 3D product design software. Autodesk Inventor was used for this design. Open Autodesk Inventor and create a new standard part.Save the file with appropriate description. Create the sketch of RHIT on the front plane using the create 2D sketch feature. Click on the rectangle icon and place its bottom left corner at the origin of the sketch plane.Draw a second rectangle inside the first one to represent the imaging window of RHIT. Add dimensions to both rectangles using the dimension feature, ensuring all lengths and widths are referenced to the origin and relative to each other. Adjust the RHIT dimensions according to the mouse size.Add thickness to RHIT by selecting outer rectangle only and extrude the sketch using the extrude tool to leave an opening in the center. Select the bottom face of the extruded RHIT and start a new 2D sketch using the start 2D sketch feature. Draw a rectangle to define the pocket for the heat pad.Define the dimensions of the rectangle based on the size and thickness of the heat pad that will be inserted into this slot. These measurements should match the physical heat pad to be used to ensure accurate temperature modeling of the mammal. Apply the required dimensions using the dimension feature.Exit the 2D sketch and perform a material removal using the cut extrude tool to create a deep cavity for the heat pad. Zoom in as needed to precisely position the sketch. Start a new sketch for the temperature probe on the same face as the cavity.Click on start a 2D sketch and select the circle icon. Place the circle on the plain surface and dimension accordingly. Exit the sketch mode.Click on extrude and select the sketch of the circle. In extrusion mode, click on cut, extrude, and remove material. Place the circle on the right side of the face with the pocket insert.It doesn't need to be centered yet. Add dimensions to align it with the center of the thickness of RHIT. Click extrude.A popup will appear with an arrow showing the direction of the cut, along with the through all option. Select through all to cut through the entire material, then click okay. Exit extrusion mode.Add fillets on the edges of window by clicking on fillet icon and select all the eight sharp edges. Add fillet size and exit fillet mode. Prepare the model for 3D printing by navigating to the environment tab and selecting 3D print tab.To print the unmodified RHIT, import the STL file, either from figure 1A or the supplementary materials, into A 3D printing preparation software. Click import, select the file, and click prepare to slice the model into printable layers. Choose a compatible commercial 3D printer and click print.Monitor the printing process. Once it's complete, carefully remove and clean the printed RHIT as needed. Wait 10 to 15 minutes after the ejections before beginning surgery and imaging.Microscope preparation and utilizing RHIT. Prepare a conventional inverted confocal microscope for liver IBN, as shown in figure 2A. Insert the stage adapter to accommodate the sample.For this protocol, a custom made metal insert was used with a 35 millimeter circular opening and a 40 millimeter cover glass. Thoroughly clean both sides of the cover glass before and after imaging, using 70%ethanol as necessary. Secure the appropriate RHIT based on mouse size with tape by aligning the RHIT aperture with objective lens, parallel to the imaging covering slip, as shown in figure 2G, on the metal insert with tape over the cover glass.Select the objective for imaging based on an on-area size, spatial resolution, and imaging time, such as a 25X per 95 water objective. Level the objective close to the cover glass. Secure a strip of gauze across the wet window with tape.This will help position the tissue and separate it from other abdominal organs. Prewarm RHIT using a heat pad inserted into the slot and secure the heat probe adjacent to the RHIT, shown in figure 2G through H.This will maintain the body temperature of the mouse during imaging. Shave the abdominal coat using a hair clipper and/or hair removal cream.Move hair residue and disinfect the exposed skin with 70%ethanol. Depending on institutional IACUC requirements, an alternating scrub of Betadine and ethanol may be used to ensure appropriate surgical site preparation. Reassess the depth of anesthesia prior to surgery on the animal.Expose the liver by making a horizontal crescent-shaped two to three centimeter incision under the ribcage with surgical scissors from the xiphoid, ending at the left lateral side of the upper abdomen. After the initial incision, carefully make a smaller secondary incision in the muscle layer to expose the left lobe of the liver. Cauterize as necessary to avoid bleeding.Prevent heat damage to the liver while using the cauterizer by placing a saline-soaked gauze between the liver and muscle layer to absorb the emanating heat of the muscle. Avoid any excessive bleeding, surgical or mechanical trauma, or external compression. Once the liver is exposed, transfer the mouse to the microscope stage.Positioning of the mouse on the microscope stage. Position the anesthetized mouse's abdomen facing down on top of the RHIT platform of the microscope. Adjust the nose cone so that the mouse remains under anesthesia.Use a saline-soaked cotton-tipped applicator to gently guide the left lateral lobe of the liver against the cover slip while gently applying external pressure to the chest cavity. Take advantage of gravity to carefully maneuver the liver out and onto the cover glass. Use a gauze strip to gently separate the liver from the mouse's body.Check the nose cone again to ensure the mouse remains under anesthesia during imaging. Prevent tissue dehydration by applying a water-based gel around the edges of the exposed cavity and avoiding direct contact with the heat pad. A microscope with an enclosed humidity chamber can also help maintain temperature and humidity.Apply opthalmic ointment to the eyes to prevent corneal drying. Lastly, set up timelapse IVM to begin imaging. Open laser shutters, evaluate fluorescent labeling, and identify a stable region of interest with a notable blood flow.Select appropriate image acquisitions, including format, speed, imaging intervals, and length of time, frame average, et cetera. Define upper and lower boundary for Z-stacks if required, and start timelapse recording. To validate our protocol, we quantified image stability using the structural similarity index, or SIM, shown in figure 3D.SIM compares two images based on luminance, contrast, and structural patterns. A value of one indicates perfect similarity, while zero reflects distortion. Figure 3C shows that RHIT consistently yields the highest SIM values, up to 50 times greater than no insert, demonstrating superior image quality.In images labeled A through C, we compared liver IVM without a stabilizer, with the cardboard inset, and with RHIT. RHIT maintains clear resolution of lipid droplets in magenta, nuclei in green, and vasculature in cyan. Black and white insets highlight image stability.RHIT also enabled long-term IBM in obese mice for over eight hours, as shown in movie one. Together, these results confirmed that RHIT significantly improves imaging quality and reproducibility across conditions. Taken together, RHIT simplifies the complex setup traditionally required for hepatic IBM, improving both reproducibility and accessibility.This approach anticipates a broader movement toward leveraging 3D printing for the standardization of advanced imaging protocols. Beyond liver imaging, RHIT's design principles may be extended to other applications, including organ tomography, material science, and engineering, where motion stability is critical for image fidelity.

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Research

JoVE Journal

Engineering

A method of visualizing and quantifying the 3-dimensional structure of mouse hepatic portal vein or intrahepatic bile duct is described. This resin cast technique can also be applied to other ductal or vascular systems and allows for in situ visualization or quantification of a system's intact communicating architecture.

3-Dimensional Resin Casting and Imaging of Mouse Portal Vein or Intrahepatic Bile Duct System

This video describes an imaging procedure to observe and study intravascular dynamics of a mouse liver tissue using multiphoton microscopy following stabilization of the organ. Multiphoton microscopy enables the imaging of biological tissue at high spatial resolution in vivo.

Imaging Fixture Supported Mouse Liver Staging for Multiphoton Microscopy Based Vascular Visualization: A Stabilization Device-assisted Imaging Technique to Obtain Multi-dimensional Images of Blood Vessels in Mouse Liver

Stable intravital high-resolution imaging of immune cells in the liver is challenging. Here we provide a highly sensitive and reliable method to study migration and cell-cell-interactions of immune cells in mouse liver over long periods (about 6 hours) by intravital multiphoton laser scanning microscopy in combination with intensive care monitoring.

Long Term Intravital Multiphoton Microscopy Imaging of Immune Cells in Healthy and Diseased Liver Using CXCR6.Gfp Reporter Mice

Double resin casting micro-computed tomography, or DUCT, enables visualization, digitalization, and segmentation of two tubular systems simultaneously to facilitate 3D analysis of organ architecture. DUCT combines ex vivo injection of two radiopaque resins followed by micro-computed tomography scanning and segmentation of the tomographic data.

DUCT: Double Resin Casting followed by Micro-Computed Tomography for 3D Liver Analysis

The present protocol optimizes the liver in situ perfusion/decellularization and two-photon microscopy methods to establish a reliable platform to visualize the dynamics of extracellular matrix (ECM) remodeling during non-alcoholic steatohepatitis (NASH).

3D Imaging of the Liver Extracellular Matrix in a Mouse Model of Non-Alcoholic Steatohepatitis

Using a diet-induced non-alcoholic fatty liver disease (NAFLD) mouse model, we describe the use of novel in vivo micro-computed tomography imaging techniques as a non-invasive method to assess the progression stages of NAFLD, focusing predominantly on the hepatic vascular network due to its significant involvement in NAFLD-related hepatic dysregulation.

Novel In Vivo Micro-Computed Tomography Imaging Techniques for Assessing the Progression of Non-Alcoholic Fatty Liver Disease

The objective of this study was to develop a novel three-dimensional digital model for the early diagnosis of hepatic fibrosis, which includes the stiffness of each voxel in the patient's liver and can thus, be used to calculate the distribution ratio of the patient's liver at different fibrosis stages.

A Three-Dimensional Digital Model for Early Diagnosis of Hepatic Fibrosis Based on Magnetic Resonance Elastography

The protocol presents a new tool to simplify intravital imaging using inverted confocal microscopy.

Streamlined Intravital Imaging Approach for Long-Term Monitoring of Epithelial Tissue Dynamics on an Inverted Confocal Microscope

Here, we describe a platform that allows noninvasive in vivo imaging of liver spheroids engrafted in the anterior chamber of the mouse eye. The workflow spans from generating spheroids from primary liver cells to transplantation into the mouse eye and in vivo imaging at cellular resolution by confocal microscopy.

In Vivo Imaging of Liver Spheroids Engrafted in the Anterior Chamber of the Mouse Eye

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.

Mapping Hepatic Stellate Cell Morphology in Mouse Models of Liver Fibrosis

Learn how to fabricate and use the Reusable Hepatic Imaging Tool for high-resolution intravital microscopy of the liver in live mice. This 3D-printed device improves tissue stability and reduces motion artifacts during extended imaging sessions.

Reusable Hepatic Imaging Tool: A Reusable 3D-Printed Tool for Hepatic Intravital Microscopy

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