This work was supported by the Swedish Diabetes Association, Funds of Karolinska Institutet, The Swedish Research Council, Novo Nordisk Foundation, The Family Erling-Persson Foundation, Strategic Research Program in Diabetes at Karolinska Institutet, The Family Knut and Alice Wallenberg Foundation, The Jonas &#38; Christina af Jochnick Foundation, Swedish Association for Diabetology and ERC-2018-AdG 834860-EYELETS. The figure drawings were created by FL-B using BioRender.com.

All procedures performed on animals were approved by the Animal Experiment Ethics Committee at Karolinska Institutet.
1. Extraction of primary mouse liver cells and generation of liver spheroids&#160;&#160;in vitro
<ol>
	<li>Preparation
	<ol>
		<li>For cannulation, liver resection, and isolation of primary liver cells, prepare the following sterile or single-use materials in addition to serological pipettes and centrifuge tubes (Figure 1A): peristaltic pump, swing-bucket centrifuge, absorbent pad, dissection mat, two curved-tip forceps, surgical scissors, one 27 G butterfly needle, one 70 &#181;m cell strainer, one cell lifter, one 100 mm Petri dish, cell-counting chamber, and 96-Well, U-shaped-bottom microplates.</li>
		<li>Prepare the solutions used in the liver perfusion, isolation of primary liver cells, and generation of liver spheroids as listed in Table 1. 
		NOTE: The Digestion buffer and the Gradient solution should be prepared fresh.</li>
		<li>Set up the perfusion system consisting of a peristaltic pump, which conducts solutions from the 42 &#176;C water bath to the liver (Figure 1A). The higher temperature of the water bath ensures that the buffers reach the liver at the optimal temperature of 37 &#176;C. Customize this depending on the tube length and the room temperature (RT).</li>
		<li>For the cannulation, fit a 27 G butterfly needle to the end of the tube. Keep the plating media used in the final isolation steps at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Procedure
	<ol>
		<li>Pre-heat the following solutions in the 42 &#176;C water bath in 50 mL centrifuge tubes: 40 mL of PBS, 20 mL of perfusion buffer, and 12 mL of digestion buffer.</li>
		<li>To clean and warm the pump tubes, circulate approximately 20 mL of the pre-heated PBS.</li>
		<li>Change the tubing to the perfusion buffer, prime the tube and butterfly needle, and set the flow rate to 4 mL/min. Ensure no bubbles in the tubing during buffer changes throughout the protocol.</li>
		<li>Euthanize the mouse by cervical dislocation and use needles to fix the limbs to the dissecting board.</li>
		<li>Wet the abdomen fur with 70% ethanol and dissect it to access the digestive organs.</li>
		<li>Move the intestines to the right to expose the portal vein and inferior vena cava (Figure 1B).</li>
		<li>Cannulate the vena cava at approximately halfway of its length with the needle in a horizontal position, ensure it is stable, and start the pump.</li>
		<li>When the liver starts to inflate, or white dots appear in the closer lobes, cut the portal vein to allow the blood and perfusion buffer to drain out.</li>
		<li>The liver should start blanching immediately. Encourage clearing using curved forceps to clamp the portal vein at 5 s intervals.</li>
		<li>Repeat step 1.2.9 until the liver is yellow and cleared of blood (approx. 15-20 mL of perfusion buffer).</li>
		<li>Stop the peristaltic pump to change the tubing to the digestion buffer and restart the pump. When the digestion buffer reaches the liver, reduce the flow rate to 2.5 mL/min. 
		NOTE: The phenol red in the digestion buffer allows to discriminate its arrival to the liver and enables the adjustment of pump parameters during the treatment.</li>
		<li>To encourage the buffer to reach all hepatic lobes and ensure proper digestion, repeat step 1.2.9 several times.</li>
		<li>Stop the flow of the peristaltic pump when the digestion buffer is depleted or the liver looks sufficiently digested. 
		NOTE: The degree of digestion can be visually monitored by gently pinching the liver lobes with forceps and checking if small marks appear on the tissue. The liver will also become flimsy.</li>
		<li>To extract the liver, cut the hepatic ligaments and connections within the abdominal cavity, aiming to remove it entirely, and place it in the Petri dish containing 10 mL of cold plating medium (Table 1).</li>
		<li>After removing the gallbladder, make small pinches on the lobes using forceps, lightly tearing the hepatic capsule. By shaking the liver in the dish, observe cells pouring out into the medium.</li>
		<li>Holding the liver steady with forceps, gently drag the cell lifter along the lobes to release the cells. 
		NOTE: Correct interlobular digestion will lead to cell suspension in the media, not tissue fragments.</li>
		<li>Using a serological pipette, collect the cell suspension from the Petri dish and filter through the 70 &#181;m cell strainer placed on a 50 mL centrifuge tube. Use fresh plating media to wash the dish of digested liver cells and transfer them to the filter.</li>
		<li>Centrifuge at 50 x g for 5 min at 4 &#176;C to pellet the cells.</li>
		<li>Remove the supernatant, leaving approximately 1 mL to cover the cell pellet, swirl the tube to resuspend the cells, and then gradually add 10 mL of cold plating medium.</li>
		<li>Add the 10 mL of gradient solution to the cell suspension and gently invert the tube 10 times.</li>
		<li>Centrifuge at 200 x g for 10 min at 4 &#176;C.</li>
		<li>The pellet contains viable liver cells enriched for hepatocytes, while the supernatant contains dead cells and debris. Discard the supernatant using a serological pipette, leaving approximately 1 mL, and resuspend the pellet by gentle swirling.</li>
		<li>Add 20 mL of cold plating medium to the cell suspension and centrifuge at 50 x g for 5 min at 4 &#176;C to wash off the gradient solution.</li>
		<li>Remove the supernatant, leaving approximately 1 mL above the pellet, and resuspend the cells in 20 mL of cold plating medium. 
		NOTE: Here, the cell pellet can be compacted, so if needed, use a 10 mL serological pipette to gently dissociate the cells.</li>
		<li>Manually determine the cell number and viability using a cell counting chamber and Trypan Blue. 
		NOTE: The hepatocyte cells will quickly precipitate in the tube; to resuspend them gently invert the tube a few times.</li>
		<li>Seed the liver cells in 200 &#181;L/well of plating medium at 1200 cells/well into 96-well ultra-low adherence plates. 
		NOTE: The optimal volume of media per well is 200 &#181;L; however, it is possible to seed cells in 100 &#181;L/well.</li>
		<li>Spin the plates at 200 x g for 3 min to gather the cells in the center of the wells.</li>
		<li>Incubate (37 &#176;C, 5% CO2) the cells and leave them to form spheroids for 5 days (Figure 1C) naturally.</li>
		<li>On day 5, carefully remove half of the media in the well and replace it with serum-free maintenance media (Table 1). Repeat this step every 48 h until day 10, when the liver spheroids are ready to be transplanted. 
		NOTE: The formation of a capsule-like structure in cultured liver spheroids shows good aggregation and viability.</li>
	</ol>
	</li>
</ol>
2. Transplantation of liver spheroids into the anterior chamber of the eye (ACE)
<ol>
	<li>Preparation
	<ol>
		<li>For transplantation of liver spheroids into the ACE, ensure to arrange the following resources (Figure 2A): stereomicroscope, isoflurane anesthesia unit, induction chamber, isoflurane, heating pad, custom-made metal base plate, mouse head-holder and gas mask, forceps attached to the solid universal joint, Hamilton 500 &#181;L threaded plunger syringe, silicone, polyethylene and pump tubing, custom-made blunt glass cannula or 24 G catheter, ethanol 70%, sterile saline, sterile 23 G needles, eye ointment (liquid paraffin and Vaseline at 1:1 ratio), disposable 1 mL syringe, and cell suspension dish 35 mm.</li>
		<li>Clean the Hamilton syringe, tubing, and cannula by passing 70% ethanol and saline.</li>
		<li>Fill the Hamilton syringe, tubing, and glass cannula with saline and fix the Hamilton syringe to the bench in a horizontal position with tape (Figure 2A).</li>
		<li>Use a custom-made blunt glass cannula.
		<ol>
			<li>Elongate a borosilicate glass capillary using a dual-stage micropipette puller to an inner diameter of &#62;300 &#181;m to allow the aspiration of the spheroids.</li>
			<li>Bevel and blunt the tip using a microelectrode beveler and expose the cannula tip to a flame for a few seconds to soften the edges. 
			NOTE: The microelectrode beveler consists of a rotating sanding stone that is operated manually; therefore, specific settings are non-applicable.</li>
		</ol>
		</li>
		<li>Alternatively, construct a cannula using the plastic portion of a 24 G catheter (Figure 2B).</li>
		<li>Prepare the anesthesia isoflurane unit and warm the heating pad to 37 &#176;C.</li>
		<li>Cover the tips of the forceps attached to the solid universal joint with a piece of polyethylene tubing to form a loop, which helps to stabilize the eye.</li>
		<li>Transfer the liver spheroids from the 96-well plate into a 35 mm cell suspension dish with maintenance media using a pipette and 200 &#181;L tip.</li>
	</ol>
	</li>
	<li>Procedure
	<ol>
		<li>Anesthetize the mouse in the induction chamber using a dose of 2.5% isoflurane and 280 mL/min air.</li>
		<li>When the mouse is unconscious, lower the anesthesia to 1.8% isoflurane and 280 mL/min air, connect the anesthesia tube to the head-holder, and quickly transfer the animal onto the heating pad, positioning the nose inside the head-holder.</li>
		<li>Immobilize the head with the screws, gently pop the eye out of the socket and secure it with the forceps and place a drop of saline on both eyes to prevent drying.</li>
		<li>Under the stereoscope, use the Hamilton syringe to aspirate and collect the liver spheroids into the tip of the cannula and leave them to rest horizontally on a clean surface. 
		NOTE: Aspirating media along with the liver spheroids helps to prevent them from sticking to the cannula walls.</li>
		<li>Carefully puncture the cornea using a 23 G needle and dry the seeping aqueous humor with a tissue. If needed, to make the incision wider, carefully slide the needle sideways to slice the cornea. 
		NOTE: A single-use sterile needle is used to perform the corneal puncture, so the cornea is not disinfected prior to incision.</li>
		<li>Add saline drops to the eye to avoid drying.</li>
		<li>Take the cannula containing the liver spheroids and hold it vertically to allow the spheroids to gravitate toward the tip of the cannula.</li>
		<li>Gently insert the cannula into the hole, and with the bevel directed towards the pupil, use the Hamilton syringe to slowly expulse the liver spheroids into the ACE (Figure 2C). 
		NOTE: Before removing the cannula, waiting a few seconds for the liquid pressures inside and outside the eye to readjust and avoid the spheroids escaping back out of the eye is recommended.</li>
		<li>From outside the cornea, position the liver spheroids around the pupil and away from the incision by gently prodding the cornea with the tip of the cannula (Figure 2C).</li>
		<li>Wait ~5-10 min for the liver spheroids to settle onto the iris before releasing the eye from the forceps.</li>
		<li>Apply vaseline eye ointment to the operated eye, which helps lubricate and heal the cornea.</li>
		<li>If desired, proceed to operate on the second eye following the same method.</li>
		<li>Before awakening the mouse, administer an analgesic to avoid post-operative discomfort, for example, 0.1 mg/kg buprenorphine in sterile saline, administered subcutaneously. 
		NOTE: Only one dose of analgesics was administered to the mice as they recovered quickly from this small procedure and did not show any signs of pain or altered behavior. Since this procedure is very quick (taking under 10 min) and causes only minor discomfort, the mice require no post-operative care, other than the post-operative analgesia administered before awakening the animal.</li>
	</ol>
	</li>
</ol>
3. In vivo imaging of engrafted liver spheroids in the ACE
<ol>
	<li>Preparation
	<ol>
		<li>Prepare the following materials and instruments for noninvasive in vivo imaging of liver spheroids engrafted in ACE (Figure 3A): upright confocal microscope, long working-distance water-dipping objective, isoflurane anesthesia unit, induction chamber, isoflurane, heating pad, custom-made metal base plate, mouse head-holder and gas mask, forceps attached to the solid universal joint, artificial-tear gel, eye ointment (liquid paraffin and Vaseline at 1:1 ratio).</li>
		<li>Optional materials include injectable fluorescent probes, disposable syringes, and 27 G needles for tail intravenous injection.</li>
	</ol>
	</li>
	<li>Procedure
	<ol>
		<li>Anesthetize the mouse in the induction chamber using a dose of 2.5% isoflurane and 280 mL/min air.</li>
		<li>When the mouse is unconscious, lower the anesthesia to 1.8% isoflurane and 280 mL/min air, connect the anesthesia tube to the head-holder, and quickly transfer the animal onto the heating pad, positioning the nose inside the head-holder.</li>
		<li>Immobilize the head in the head-holder by using the screws.</li>
		<li>Place a drop of artificial tear gel on both eyes to prevent drying.</li>
		<li>At this point, intravenously inject fluorescent probes through the tail vein and image immediately after.</li>
		<li>Tilt the head, gently pop the eye out of the socket, and secure it with forceps and into position under the objective.</li>
		<li>Apply a generous amount of artificial tear gel to fill the space between the cornea and the objective and focus on the liver spheroids through the eyepiece. 
		NOTE: If possible, remove the eyepiece of one of the oculars to gain non-magnified vision and more easily locate the spheroids on the iris.</li>
		<li>In order to obtain high-resolution imaging despite the breathing movements of the animal, use 25x objectives and the following imaging settings: format 512 x 512 pixels, scan speed of 600 Hz, and a Z-stack thickness of 3 &#181;m. See detailed imaging settings in Table 2. 
		NOTE: Throughout the imaging, the anesthesia concentration is adjusted from 1.6-2.2 mL/h isoflurane to achieve a shallow, controlled breathing rhythm and thereby minimize the movement of the animal.</li>
		<li>At the end of the imaging session, treat the imaged eyes with vaseline eye ointment before removing isoflurane and awakening the animal.</li>
	</ol>
	</li>
</ol>

Noninvasive Intraocular In Vivo Imaging of Engrafted Liver Spheroids in Mice

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	<li>Bell, C. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+primary+human+hepatocyte+spheroids+as+a+model+system+for+drug-induced+liver+injury,+liver+function+and+disease.">Characterization of primary human hepatocyte spheroids as a model system for drug-induced liver injury, liver function and disease.</a> Sci Rep. 6, 25187 (2016).</li><li>Lauschke, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Massive+rearrangements+of+cellular+MicroRNA+signatures+are+key+drivers+of+hepatocyte+dedifferentiation.">Massive rearrangements of cellular MicroRNA signatures are key drivers of hepatocyte dedifferentiation.</a> Hepatology. 64 (5), 1743-1756 (2016).</li><li>Oliva-Vilarnau, N., Vorrink, S. U., Ingelman-Sundberg, M., Lauschke, V. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+3D+cell+culture+model+identifies+Wnt/beta-catenin+mediated+inhibition+of+p53+as+a+critical+step+during+human+hepatocyte+regeneration.">A 3D cell culture model identifies Wnt/beta-catenin mediated inhibition of p53 as a critical step during human hepatocyte regeneration.</a> Adv Sci (Weinh). 7 (15), 2000248 (2020).</li><li>Hurrell, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+liver+spheroids+as+a+model+to+study+aetiology+and+treatment+of+hepatic+fibrosis.">Human liver spheroids as a model to study aetiology and treatment of hepatic fibrosis.</a> Cells. 9 (4), 964 (2020).</li><li>Kozyra, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+hepatic+3D+spheroids+as+a+model+for+steatosis+and+insulin+resistance.">Human hepatic 3D spheroids as a model for steatosis and insulin resistance.</a> Sci Rep. 8 (1), 14297 (2018).</li><li>Lauschke, V. M., Shafagh, R. Z., Hendriks, D. F. G., Ingelman-Sundberg, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+primary+hepatocyte+culture+systems+for+analyses+of+liver+diseases,+drug+metabolism,+and+toxicity:+Emerging+culture+paradigms+and+applications.">3D primary hepatocyte culture systems for analyses of liver diseases, drug metabolism, and toxicity: Emerging culture paradigms and applications.</a> Biotechnol J. 14 (7), e1800347 (2019).</li><li>Shibuya, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+efficacy+of+the+hepatocyte+spheroids+for+hepatocyte+transplantation.">The efficacy of the hepatocyte spheroids for hepatocyte transplantation.</a> Cell Transplant. 30, 9636897211000014 (2021).</li><li>Hamazaki, K., Doi, Y., Koide, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microencapsulated+multicellular+spheroid+of+rat+hepatocytes+transplanted+intraperitoneally+after+90%+hepatectomy.">Microencapsulated multicellular spheroid of rat hepatocytes transplanted intraperitoneally after 90% hepatectomy.</a> Hepatogastroenterology. 49 (48), 1514-1516 (2002).</li><li>Marques, P. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+liver+biology+in+vivo+using+conventional+confocal+microscopy.">Imaging liver biology in vivo using conventional confocal microscopy.</a> Nat Protoc. 10 (2), 258-268 (2015).</li><li>Lazzeri-Barcelo, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraocular+liver+spheroids+for+noninvasive+high-resolution+in+vivo+monitoring+of+liver+cell+function.">Intraocular liver spheroids for noninvasive high-resolution in vivo monitoring of liver cell function.</a> Nat Commun. 15 (1), 767 (2024).</li><li>Speier, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noninvasive+in+vivo+imaging+of+pancreatic+islet+cell+biology.">Noninvasive in vivo imaging of pancreatic islet cell biology.</a> Nat Med. 14 (5), 574-578 (2008).</li><li>Leibiger, I. B., Berggren, P. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraocular+in+vivo+imaging+of+pancreatic+islet+cell+physiology/pathology.">Intraocular in vivo imaging of pancreatic islet cell physiology/pathology.</a> Mol Metab. 6 (9), 1002-1009 (2017).</li><li>Charni-Natan, M., Goldstein, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protocol+for+Primary+Mouse+Hepatocyte+Isolation.">Protocol for Primary Mouse Hepatocyte Isolation.</a> STAR protocols. 1 (2), 100086 (2020).</li><li>Baze, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+spheroid+primary+human+hepatocytes+in+monoculture+and+coculture+with+nonparenchymal+cells.">Three-dimensional spheroid primary human hepatocytes in monoculture and coculture with nonparenchymal cells.</a> Tissue Eng Part C Methods. 24 (9), 534-545 (2018).</li><li>Mohar, I., Brempelis, K. J., Murray, S. A., Ebrahimkhani, M. R., Crispe, I. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+nonparenchymal+cells+from+the+mouse+liver.">Isolation of nonparenchymal cells from the mouse liver.</a> Methods Mol Biol. 1325, 3-17 (2015).</li></ol>

P-OB is the cofounder and CEO of Biocrine AB, IBL, and BL are consultants for Biocrine AB.

This protocol describes a novel platform for intraocular in vivo imaging of liver spheroids engrafted in the ACE. The ACE has previously been used as a transplantation site of other organ-derived microtissues, such as pancreatic islets11,12, due to its unique engraftment microenvironment, being rich in vessels, nerves, and oxygen, and the access to imaging through the cornea. While intravital liver imaging enables the visualization of cells and processes in situ, longitudinal monitoring is not possible. Liver imaging through an abdominal window entails complex surgery, and the movement of the organ within the body makes single-cell tracking over time difficult. Hence, this novel imaging method enables noninvasive, longitudinal monitoring of liver cells at single-cell resolution.
This protocol is divided into three parts. First is the isolation of primary hepatocytes via two-step collagenase perfusion, adapted from Charni-Natan et al.13, with the difference that we perform the liver perfusion on the dead mouse instead of the anesthetized live animals. This variation brings certain advantages, such as fewer ethical considerations and the avoidance of anesthesia residue in the organism. In this work, we generate liver spheroids from the hepatocyte-enriched fraction of the isolation, but this does not exclude the potential to isolate other nonparenchymal cell populations using other specialized protocols to make coculture spheroids of diverse composition14,15.
The second part of this protocol involves the transplantation of the liver spheroids into the ACE of recipient mice. This is a quick (under 10 min) and simple surgery performed in anesthetized mice and does not require any post-operative treatment. The cornea puncture self-seals and heals over 3-5 days. Occasionally, during the healing process, some hazing is observed around the incision, but this clears within a few days. We have not experienced cases of anterior synechia in the eyes of operated animals. We perform the transplantation procedures in a clean but open-air laboratory and without issues with infections in the operated eyes. The inoculation and engraftment of spheroids in the eye do not compromise the vision or alter the behavior of the recipient animal. In this protocol, we use isoflurane anesthesia for both transplantation surgery and in vivo imaging, which is well tolerated in mice. Due to its dose-dependent effect, it can be easily adjusted throughout the procedures and brings the advantage of reducing sleeping and awakening times. However, alternative injectable anesthetics can be used. After transplantation, we generally allow 1 month for the spheroids to fully engraft, become vascularized, and innervated, before performing treatment interventions and in vivo imaging. We have also shown that transplantation and engraftment is possible using human liver spheroids and immunocompromised recipient mice10.
The third part of this method is the in vivo imaging of the engrafted liver spheroids in the ACE. This protocol describes the in vivo imaging setup, which uses microscopy equipment commonly found in research imaging facilities. Moreover, the specialized materials, such as the mouse head-holder and the plastic cannula, are now commercially available. With this imaging setup, we are able to capture z-sections and obtain a three-dimensional reconstruction of the spheroid architecture, depending on the depth of laser penetration and fluorescence detection. Monitoring cellular function in the engrafted liver spheroids relies on the visualization of fluorescent proteins that report on cell types, cellular functions, and dynamics. Thus, this imaging platform can be exploited using different modalities, alone or in combination: (1) Fluorescent probes can be administered intravenously, e.g., antibodies to label and track cells as well as functional dyes; (2) Liver spheroids can be generated from cells isolated from reporter mouse models that express liver-specific fluorescent proteins, e.g., FUCCI liver spheroids that report on cell cycle dynamics; (3) The formation of liver spheroids in vitro can be combined with transfection or transduction, to equip the spheroids with fluorescent proteins and biosensors. e.g., adeno-associated viruses. In our experimental settings and by using a single photon for excitation, the imaging depth that is possible to achieve is roughly 60-100 &#181;m. However, this is dependent on the laser power and multiphoton imaging availability, fluorescence probe emission characteristics and sensitivity of the detectors, as well as the angle of the eye in which the spheroid is engrafted. After imaging acquisition, the downstream image analysis can be performed using popular programs such as Image J and Imaris. For example, in the case of the FUCCI reporter, cell-cycle active cells in green can be counted and contrasted to the number of total red cells to assess cell-cycle activity within the engrafted spheroid. Additionally, the ACE-imaging platform allows substances to be applied to the eye (in the form of eye drops) or injected directly into the ACE to treat the graft and monitor its reaction. Post-mortem, the transplanted spheroids can easily be retrieved by manual microdissection and can provide valuable information by ex vivo techniques, such as immunofluorescence staining, transcriptomic analysis, etc.10.
This technique has certain limitations. The first is that from our experience, the recipient mice must be albino, i.e., have non-pigmented iris. Upon engraftment, the liver spheroids become covered by a monolayer of iris cells, which does not affect the viability or function of the spheroids, but the pigment in the iris cells prevents imaging. A second consideration is the stability during intraocular imaging in anesthetized mice. During the in vivo imaging sessions, the anesthesia concentration and breathing of the animal must be closely monitored to minimize movement. Nevertheless, using the imaging settings indicated here, we are able to achieve high-resolution imaging at the single-cell level.
To summarize, this protocol describes the implementation of a noninvasive in vivo imaging platform of liver-like tissue engrafted in the eyes of mice. We use easy procedures, common equipment, and affordable materials, making it an attainable approach for many investigators. This model combines the advantages of in vitro 3D-liver spheroids with the in vivo milieu and optical accessibility provided by the ACE to create a valuable platform for studying liver physiology and pathology in basic research and pre-clinical settings.

The monitoring of liver function in mammals during health and disease is limited by the lack of high-resolution, noninvasive in vivo imaging techniques. The visualization of this organ is hindered by its inaccessible location, and in order to piece together cellular processes, in vivo studies rely on the sacrifice of animals at different time points. To circumvent this imaging limitation, much work relies on in vitro models, in which liver-like microtissues are visualized and studied in a controlled environment.
In recent years, the development of three-dimensional culture systems, such as liver spheroids, has aided and advanced liver research. Liver spheroids are multicellular aggregates that mimic the microenvironment and complex cell-cell interactions of liver tissue to a certain extent1 and offer clear advantages over traditional monolayer cultures2,3. Liver spheroids are also used as models for different liver diseases4,5,6 and have been instrumental to&#160;understanding disease mechanisms. Still, key limitations of the current in vitro liver models are the lack of a physiological in vivo milieu and the limited utilization time in culture (around 20 days)3. Liver spheroids have been previously transplanted to different sites in vivo, such as under the kidney capsule7 or intraperitoneally8, which are not accessible for optical imaging. Intravital liver imaging is a state-of-the-art technique offering cell-resolution real-time imaging. Currently, this in situ liver imaging is only possible on the exteriorized organ, which is highly invasive and often terminal9. Although the fitting of an abdominal window would allow repeated liver imaging sessions, it entails complex surgery and after-care.
To perform longitudinal monitoring at cellular resolution, we explored transplantation of liver spheroids into the anterior chamber of the eye (ACE) of mice, where the liver-like tissue is engrafted in a physiological milieu, connected to the body stimuli, and accessible for optical imaging. The cornea is a transparent tissue and acts as a window through which microtissues engrafted on the iris can be imaged non-invasively and longitudinally by confocal microscopy. Here, we present a workflow of this newly developed platform for in vivo imaging of liver spheroids10. This protocol is a step-by-step guide for its implementation, divided into (1) the extraction of primary mouse liver cells and in vitro formation of liver spheroids, (2) the transplantation of liver spheroids into the ACE of recipient mice, and (3) the in vivo imaging of engrafted liver spheroids in anesthetized mice. Furthermore, we will showcase some of the possibilities and applications of this imaging platform.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>27 G butterfly needle</td>
			<td>Venofix</td>
			<td>4056388</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV8-CAG-GFP</td>
			<td>Charles River</td>
			<td>CV17169-AV9</td>
			<td>Incubated with isolated hepatocytes at 1 &#181;L/mL during liver spheroid formation</td>
		</tr>
		<tr>
			<td>Absolute and 70% ethanol</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorbent pad</td>
			<td>Attends</td>
			<td>203903</td>
			<td></td>
		</tr>
		<tr>
			<td>Albumin-Cre;RCL-tdTomato (B6.Cg-Speer6-ps1Tg(Alb-cre)21Mgn/J ; B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)</td>
			<td>Jackson</td>
			<td>#003574 and #007914</td>
			<td>Mice obtained from in-house breeding</td>
		</tr>
		<tr>
			<td>B6 albino mice (B6(Cg)-Tyrc-2J/J)</td>
			<td>Jackson</td>
			<td>#000058</td>
			<td>Mice obtained from in-house breeding</td>
		</tr>
		<tr>
			<td>B6;129P2-Gt(ROSA)26Sor[tm1(CAG-Venus/GMNN,-Cherry/CDT1)Jkn]/JknH</td>
			<td>INFRAFRONTIER/EMMA</td>
			<td>EM:08395</td>
			<td>Mice obtained from in-house breeding</td>
		</tr>
		<tr>
			<td>BD Insyte IV Catheter 24 G x 0.75 in</td>
			<td>BD Medical</td>
			<td>381212</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosillicate standard glass cappilaries&#160;</td>
			<td>World Precision Instruments</td>
			<td>1B150-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell lifter</td>
			<td>Corning</td>
			<td>3008</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer, 70 &#181;m</td>
			<td>Falcon</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-made metal plate</td>
			<td>Hardware store</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Dexamethasone</td>
			<td>Sigma-Aldrich</td>
			<td>D4902</td>
			<td></td>
		</tr>
		<tr>
			<td>Dual-Stage Glass Micropipette Puller</td>
			<td>Narshige</td>
			<td>Model PC-100</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>E9884</td>
			<td></td>
		</tr>
		<tr>
			<td>Electric heating pad</td>
			<td>Hardware store</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Gibco</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMAX</td>
			<td>Gibco</td>
			<td>35050061</td>
			<td></td>
		</tr>
		<tr>
			<td>Green CMFDA</td>
			<td>Abcam</td>
			<td>ab145459</td>
			<td>Reconstituted in DMSO, administered at 100 &#181;g/mouse in PBS 10% FBS</td>
		</tr>
		<tr>
			<td>Hamilton syringe</td>
			<td>Hamilton</td>
			<td>81242</td>
			<td>Model 1750 Luer Tip Threaded Plunger Syringe, 500 &#181;L</td>
		</tr>
		<tr>
			<td>HBSS; no calcium, no magnesium and no phenol red</td>
			<td>Gibco</td>
			<td>14175095</td>
			<td></td>
		</tr>
		<tr>
			<td>HCX IRAPO L 25x/0.95 W objective</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Gibco</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>Induction chamber 0.8 L</td>
			<td>Univentor</td>
			<td>8329001</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin-Transferrin-Selenium (ITS-G)</td>
			<td>Gibco</td>
			<td>41400045</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Baxter</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Lectin DyLight-649</td>
			<td>Invitrogen</td>
			<td>L32472</td>
			<td>Administered at 1 mg/mL and 100 &#181;L/mouse</td>
		</tr>
		<tr>
			<td>Liberase TM Research Grade</td>
			<td>Sigma-Aldrich</td>
			<td>5401127001</td>
			<td></td>
		</tr>
		<tr>
			<td>Microelectrode beveler</td>
			<td>World Precision Instruments</td>
			<td>Model BV-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse head-holder and gas mask</td>
			<td>Narshige</td>
			<td>Model SGM-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunclon Sphera 96-Well, U-Shaped-Bottom Microplate</td>
			<td>Thermo Fisher</td>
			<td>174929</td>
			<td></td>
		</tr>
		<tr>
			<td>Oculentum simplex</td>
			<td>APL</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS 10x</td>
			<td>Gibco</td>
			<td>14080055</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS 1x; no calcium, no magnesium</td>
			<td>Gibco</td>
			<td>14190144</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Percoll</td>
			<td>Sigma-Aldrich</td>
			<td>P1644</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Ismatec</td>
			<td>Model ISM795</td>
			<td></td>
		</tr>
		<tr>
			<td>PharMed BPT Pump Tubing</td>
			<td>VWR</td>
			<td>VERN070540-07</td>
			<td>Inner diameter 0.76 mm, outer diameter 2.46 mm</td>
		</tr>
		<tr>
			<td>pHrodo Red-LDL</td>
			<td>Invitrogen</td>
			<td>L34356</td>
			<td>Administered at 1 mg/mL and 100 &#181;L/mouse</td>
		</tr>
		<tr>
			<td>Portex Fine Bore Polyethylene Tubing</td>
			<td>Smiths Medical</td>
			<td>800/100/140</td>
			<td>Inner diameter 0.4 mm, outer diameter 0.8 mm</td>
		</tr>
		<tr>
			<td>Silicone dissection mat</td>
			<td>Hardware store</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride 0.9%</td>
			<td>Braun</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Solid Universal Joint</td>
			<td>Narshige</td>
			<td>Model UST-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>Model M80</td>
			<td></td>
		</tr>
		<tr>
			<td>Suspension culture dish 35 mm</td>
			<td>Sarstedt</td>
			<td>833900500</td>
			<td></td>
		</tr>
		<tr>
			<td>Temgesic</td>
			<td>Indivor</td>
			<td>N/A</td>
			<td>Administered s.c. at 0.05 mg/mL and 2 &#181;L/g mouse</td>
		</tr>
		<tr>
			<td>Translucent Silicone Tubing</td>
			<td>VWR</td>
			<td>228-1450</td>
			<td>Inner diameter 1.5 mm, outer diameter 3 mm</td>
		</tr>
		<tr>
			<td>Trypan Blue</td>
			<td>Sigma-Aldrich</td>
			<td>T8154</td>
			<td></td>
		</tr>
		<tr>
			<td>Univentor 400 Anesthesia unit&#160;</td>
			<td>Univentor</td>
			<td>8323001</td>
			<td></td>
		</tr>
		<tr>
			<td>Upright laser scanning confocal microscope</td>
			<td>Leica</td>
			<td>Model TCS SP5 II</td>
			<td></td>
		</tr>
		<tr>
			<td>Viscotears</td>
			<td>Novartis</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>William&#39;s E Medium; no glutamine, phenol red</td>
			<td>Gibco</td>
			<td>22551089</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo imaging of liver spheroids engrafted in the anterior chamber of the mouse eye

Biomedical studies of the liver in mammals are hindered by the lack of methods for in vivo noninvasive longitudinal imaging at cellular resolution. Until now, optical imaging of the liver in situ is possible by intravital imaging, which offers high-resolution imaging at the cellular level but cannot be performed multiple times and, therefore, longitudinally in the same animal. Noninvasive imaging methods, such as bioluminescence, allow repeated imaging sessions on the same animal but do not achieve cell resolution. To address this methodology gap, we have developed a platform for noninvasive in vivo imaging of liver spheroids engrafted in the anterior chamber of the mouse eye. In the workflow described in this study, primary mouse liver spheroids are generated in vitro and transplanted into the anterior chamber of the eye of recipient mice, where they engraft on the iris. The cornea acts as a natural body window through which we can image the engrafted spheroids by conventional confocal microscopy. The spheroids survive for months in the eye, during which the cells can be studied in contexts of health and disease, as well as being monitored in response to different stimuli over repeated imaging sessions using appropriate fluorescent probes. In this protocol, we provide a breakdown of the necessary steps to implement this imaging system and explain how to best harness its potential.

Primary liver cells, enriched for hepatocytes, were isolated from the mouse liver by two-step collagenase perfusion, using a peristaltic pump to circulate warm buffers through the liver taking advantage of the organ&#39;s vasculature to deliver dissociation enzymes to all cells (Figure 1A). For this, the inferior vena cava was cannulated, and the portal vein was snipped to allow the flow-through of buffers (Figure 1B). First, an HBSS-based buffer was flushed through the liver to clear the blood. If the cannulation is successful and there are no blood clots, the liver blanches and becomes yellow within a few seconds. Secondly, a Digestion buffer containing the Liberase enzyme blend was circulated through the liver to dissociate the tissue into a single-cell suspension. The cells were manually counted and seeded into 96-well ultra-low-adherence (ULA) plates, which enable the self-assembly into spheroids within a few days. On day 5, the spheroids are formed, and the thin capsule bordering the spheroids indicates successful aggregation (Figure 1C). We wait until day 10 to transplant, at which point the spheroids are compact and have developed strong cell-cell connections. The number of seeded cells per well determined the size of the liver spheroid, with 1000, 1200, and 1500 cells/well yielding spheroids of 238 &#181;m &#177; 10 &#181;m, 248 &#181;m &#177; 17 &#181;m, and 298 &#181;m &#177; 19 &#181;m (mean &#177; SD) diameters, respectively (Figure 1C,D). For transplantation, we select spheroids of approximately 250 &#181;m diameter for the following reasons: (1) the spheroids size should not be too large to avoid hypoxia and necrotic core, but should contain enough cells to support cell-cell communications and to allow graft remodeling in the eye, (2) the weight of spheroids of this size allows them to gravitate towards the iris and improve their engraftment, (3) this size is appropriate in relation to transplanting 5-10 spheroids per mouse eye.
The transplantation surgery requires a manual-threaded syringe connected to a glass cannula (Figure 2A). The glass cannula consists of a borosilicate glass capillary modified in-house to have a fine blunt tip using a micropipette puller and beveler. A simpler alternative cannula can be created using a commercially available plastic catheter connected to the syringe tubing and stabilized in a pipette tip (Figure 2B). The surgery consists of the inoculation of liver spheroids into the ACE through an incision in the cornea (Figure 2C). The spheroids were positioned on the borders of the pupil to make them better accessible for imaging and avoid them moving into the ocular angle. Albino mice were used for transplantation, as their non-pigmented iris allow the in vivo imaging of the engrafted liver spheroids. Recipient mice were transplanted into both eyes with 7-10 spheroids/eye, and stereoscopic images were taken at 3 days post-transplantation (post-Tx) as well as at 1 week and 1 month post-Tx to document the cornea healing and spheroid engraftment success (Figure 2D). Of note, the change in appearance of the liver spheroids in the ACE between when freshly transplanted and fully engrafted is due to the settlement of the graft onto the iris, as well as to the growth of a monolayer of iris cells over the spheroid. The engraftment success rate of liver spheroids in the ACE is 70% (n = 9 eyes in both male and female mice) (Figure 2E). The first days post-Tx are the most critical for survival and engraftment, likely due to the recipient animal rubbing its eyes and dislodging the spheroids before the cornea has healed. The size of the liver spheroids does not differ significantly post-Tx and changes to shape are attributed to graft remodeling and engraftment (Figure 2F). At 1 month post-Tx, all engrafted spheroids present on the iris were vascularized and innervated, as shown by immunofluorescence staining (Figure 2G).
Noninvasive in vivo imaging is performed on anesthetized recipient mice using an upright confocal microscope and long-distance dipping objective (Figure 3A, Table 2). The fluorescence imaging in the ACE can be achieved through different approaches, as depicted in Figure 3B. The injection of fluorescent probes into the circulation of the recipient mouse allows the visualization of different cell types and structures within the spheroids. We used lectin to mark blood vessels (Figure 3C), CMFDA to observe the bile canaliculi network (Figure 3D) and pHrodo-LDL, which confirmed active LDL-uptake into spheroid cells (Figure 3E). Liver spheroids generated from reporter mouse models can also be used. Albumin-Cre:tdTomato spheroids allowed labeling and tracking of hepatocytes (Figure 3F), and spheroids expressing the Fluorescent Ubiquitin Cell Cycle Indicator (FUCCI) biosensor were used to visualize cell cycle dynamics at single-cell resolution (Figure 3G). Finally, liver spheroids can be genetically modified in vitro prior to transplantation, and, in the case of adeno-associated virus (AAV)-GFP transduction, the expression was observed in vivo for over 6 months (Figure 3H).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66234/66234fig01.jpg" /> 
Figure 1: Isolation of primary mouse hepatocytes and generation of liver spheroids. (A) Material and equipment used for isolation of primary mouse hepatocytes: 1. Isolation buffers; 2. Water bath; 3. Peristaltic pump; 4. Petri dish; 5. Cell strainer; 6. Absorbent pad; 7. Dissection mat; 8. Cell Lifter; 9. Butterfly needle 27 G; 10. Dissection tools. (B) Abdominal cavity during surgery: the vena cava is cannulated and perfused, and the portal vein is snipped to allow flow-through of the buffers. (C) Brightfield images of the formation of hepatic spheroids in vitro at 0 (d0), 5 (d5), and 10 (d10) days post-seeding, scale bars = 200 &#181;m. (D) Liver spheroid size at different cell-seeding concentrations, n = 21 spheroids. <a href="https://www.jove.com/files/ftp_upload/66234/66234fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66234/66234fig02.jpg" /> 
Figure 2: Transplantation and engraftment of liver spheroids into the ACE of mice. (A) Materials and equipment used for transplantation (Tx) of liver spheroids into the ACE: 1. Liver spheroids in culture dish; 2. Sterile saline; 3. Eye ointment; 4. Needles 23 G; 5. Cannula; 6. Hamilton syringe; 7. Anesthesia gas tube; 8. Head-holder and gas mask; 9. Heating pad; 10. Custom-made metal base plate; 11. Forceps and solid universal joint. (B) Cannula and Hamilton syringe setup: 1. Glass cannula connected to the Hamilton syringe via Portex tubing and a 27G needle; 2. Glass cannula is connected to the Portex tubing via additional segments of Silicone tubing and PharMed tubing; 3. Alternative assembled plastic cannula; 4. Parts forming the plastic cannula: 24G BD Insyte plastic catheter connected via PharMed tubing and sheathed in a cut-off 10 &#181;l pipette tip for stability and grip. (C) Illustration of Tx surgery steps: 1. The spheroids are collected into the cannula; 2. The cornea is punctured with a needle; 3. The cannula is inserted into the incision, and the spheroids are released into the ACE; 4. From the outside of the eye, the spheroids are positioned close to the pupil and away from the incision. (D) Stereoscopic images of liver spheroids (sph) in the mouse eye on the day of surgery and at 3-, 7-, and 30-days post-Tx. Arrows indicate viable spheroids. (E) Liver spheroid (size of 1200 cells/well) engraftment rate post-Tx, n= 9 eyes in 6 recipient mice. (F) Size of liver spheroids in culture, prior to transplantation (in vitro, n= 20 spheroids from single preparation) and at 1-month post-Tx in the ACE (in vivo, n= 16 spheroids in 3 recipient mice), calculated by averaging vertical and horizontal diameters. (G) Immunofluorescence staining of engrafted liver spheroids at 2 months post-Tx, showing vascularization (CD31, pink, dashed line delineates the spheroid mass) and sympathetic innervation (tyrosine hydroxylase (TH), orange), scale bar = 100 &#181;m. Data for panel F has been adapted with permission from Lazzeri-Barcelo et al.10. <a href="https://www.jove.com/files/ftp_upload/66234/66234fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66234/66234fig03.jpg" /> 
Figure 3: Noninvasive intraocular in vivo imaging of engrafted liver spheroids. (A) Material and equipment used for in vivo ACE imaging: 1. Upright laser scanning confocal microscope; 2. Dark box; 3. Motorized XYZ stage; 4. Dipping-objective; 5. Head-holder and gas mask; 6. Forceps and solid universal joint; 7. Heating pad; 8. Custom-made metal base plate. (B) Diagram depicting different approaches used for in vivo imaging of fluorescent readouts in liver spheroids engrafted in the eye. (C-H) Representative images of ACE-liver spheroids during in vivo imaging by confocal microscopy. The backscatter signal is used to observe the spheroid volume and structure; (C) Blood vessels labeled by i.v. injection of fluorescent lectin, scale bar = 100 &#181;m; (D) Bile canaliculi network labeled by injection of fluorescent CMFDA, scale bar = 50 &#181;m; (E) LDL uptake by injection of fluorescent pHrodo-LDL probe, scale bar= 100 &#181;m; (F) Td-Tomato-expressing hepatocytes, arrowheads indicate nuclei and asterisks indicate intra-spheroid vasculature, scale bar = 50 &#181;m; (G) Monitoring of cell cycle dynamics in FUCCI-expressing liver spheroids, scale bars = 50 &#181;m (main image) and 20 &#181;m (blow-up). (H) liver spheroids transduced in vitro with AAV8-GFP prior to Tx and imaged in the eye at 6 months post-Tx, scale bar = 50 &#181;m. The image in panel G has been adapted with permission from Lazzeri-Barcelo et al.10. <a href="https://www.jove.com/files/ftp_upload/66234/66234fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Solutions used for the isolation of primary mouse hepatocytes. Composition of solutions and buffers needed for mouse hepatocyte isolation. The Digestion buffer and Gradient solution components should be mixed fresh on the day of isolation. <a href="https://www.jove.com/files/ftp_upload/66234/Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
Table 2. Confocal Leica SP5 microscope settings used for intraocular in vivo imaging of liver spheroids. The table has been adapted with permission from Lazzeri et al.10. <a href="https://www.jove.com/files/ftp_upload/66234/Table 2.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Author Spotlight: Harnessing Mouse Eye Chambers for Noninvasive Liver Spheroid Studies at JoVE.com

Author Spotlight: Harnessing Mouse Eye Chambers for Noninvasive Liver Spheroid Studies

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

Biomedical studies of the liver in mammals are hindered by the lack of methods for in vivo noninvasive longitudinal imaging at cellular resolution. Until ...

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1.1K Views.  Karolinska University Hospital. 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. 

Video: Author Spotlight: Harnessing Mouse Eye Chambers for Noninvasive Liver Spheroid Studies

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Do-It-Yourself Device for Recovery of Cryopreserved Samples Accidentally Dropped into Cryogenic Storage Tanks

Liquid nitrogen is colorless, odorless, extremely cold (-196 &deg;C) liquid kept under pressure. It is commonly used as a cryogenic fluid for long term storage of biological materials such as blood, cells and tissues 1,2. The cryogenic nature of liquid nitrogen, while ideal for sample preservation, can cause rapid freezing of live tissues on contact - known as 'cryogenic burn'2, which may lead to severe frostbite in persons closely involved in storage and retrieval of samples from Dewars. Additionally, as liquid nitrogen evaporates it reduces the oxygen concentration in the air and might cause asphyxia, especially in confined spaces2.
In laboratories, biological samples are often stored in cryovials or cryoboxes stacked in stainless steel racks within the Dewar tanks1. These storage racks are provided with a long shaft to prevent boxes from slipping out from the racks and into the bottom of Dewars during routine handling. All too often, however, boxes or vials with precious samples slip out and sink to the bottom of liquid nitrogen filled tank. In such cases, samples could be tediously retrieved after transferring the liquid nitrogen into a spare container or discarding it. The boxes and vials can then be relatively safely recovered from emptied Dewar. However, the cryogenic nature of liquid nitrogen and its expansion rate makes sunken sample retrieval hazardous. It is commonly recommended by Safety Offices that sample retrieval be never carried out by a single person. Another alternative is to use commercially available cool grabbers or tongs to pull out the vials3. However, limited visibility within the dark liquid filled Dewars poses a major limitation in their use.
In this article, we describe the construction of a Cryotolerant DIY retrieval device, which makes sample retrieval from Dewar containing cryogenic fluids both safe and easy.

Here we present a low cost, durable cryotolerant device for sample retrieval from Dewar tanks filled with liquid nitrogen. The ease of construction and modular design of the device makes the process of sample retrieval from cryogenic tanks safe and easy.

Liquid nitrogen is colorless, odorless, extremely cold (-196 &deg;C) liquid kept under pressure. It is commonly used as a cryogenic fluid for long term ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

A Highly Reproducible and Straightforward Method to Perform In Vivo Ocular Enucleation in the Mouse after Eye Opening

Enucleation or the surgical removal of an eye can generally be considered as a model for nerve deafferentation. It provides a valuable tool to study the different aspects of visual, cross-modal and developmental plasticity along the mammalian visual system1-4.
Here, we demonstrate an elegant and straightforward technique for the removal of one or both eyes in the mouse, which is validated in mice of 20 days old up to adults. Briefly, a disinfected curved forceps is used to clamp the optic nerve behind the eye. Subsequently, circular movements are performed to constrict the optic nerve and remove the eyeball. The advantages of this technique are high reproducibility, minimal to no bleeding, rapid post-operative recovery and a very low learning threshold for the experimenter. Hence, a large amount of animals can be manipulated and processed with minimal amount of effort. The nature of the technique may induce slight damage to the retina during the procedure. This side effect makes this method less suitable as compared to Mahajan et al. (2011)5 if the goal is to collect and analyze retinal tissue. Also, our method is limited to post-eye opening ages (mouse: P10 - 13 onwards) since the eyeball needs to be displaced from the socket without removing the eyelids. The in vivo enucleation technique described in this manuscript has recently been successfully applied with minor modifications in rats and appears useful to study the afferent visual pathway of rodents in general.

A Highly Reproducible and Straightforward Method to Perform In Vivo Ocular Enucleation in the Mouse after Eye Opening

The removal of eyes, also called enucleation, provides a useful strategy to study aspects of visual, cross-modal, and developmental plasticity along the mammalian visual system since it induces irreversible partial (monocular) or complete (binocular) vision loss. Here we describe a highly reproducible and straightforward approach to perform in vivo enucleation.

Enucleation or the surgical removal of an eye can generally be considered as a model for nerve deafferentation. It provides a valuable tool to study the ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

The Mouse Isolated Perfused Kidney Technique

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a prerequisite for studying the physiology of the isolated organ and for many innovative applications that may be possible in the future, including perfusion decellularization for kidney bioengineering or the administration of anti-rejection or genome-editing drugs in high doses to prime the kidney for transplantation. During the time of the perfusion, the kidney can be manipulated, renal function can be assessed, and various pharmaceuticals administered. After the procedure, the kidney can be transplanted or processed for molecular biology, biochemical analysis, or microscopy.
This paper describes the perfusate and the surgical technique needed for the ex vivo perfusion of mouse kidneys. Details of the perfusion apparatus are given and data are presented showing the viability of the kidney&#39;s preparation: renal blood flow, vascular resistance, and urine data as functional, transmission electron micrographs of different nephron segments as morphological readouts, and western blots of transport proteins of different nephron segments as molecular readout.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. The buffers and surgical technique are described in detail.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a ...

Real Time In Vivo Tracking of Thymocytes in the Anterior Chamber of the Eye by Laser Scanning Microscopy

The purpose of the method being presented is to show, for the first time, the transplant of newborn thymi into the anterior eye chamber of isogenic adult mice for in vivo longitudinal real-time monitoring of thymocytes&#180; dynamics within a vascularized thymus segment. Following the transplantation, laser scanning microscopy (LSM) through the cornea allows in vivo noninvasive repeated imaging at cellular resolution level. Importantly, the approach adds to previous intravital T-cell maturation imaging models the possibility for continuous progenitor cell recruitment and mature T-cell egress recordings in the same animal. Additional advantages of the system are the transparency of the grafted area, permitting macroscopic rapid monitoring of the implanted tissue, and the accessibility to the implant allowing for localized in addition to systemic treatments. The main limitation being the volume of the tissue that fits in the reduced space of the eye chamber which demands for lobe trimming. Organ integrity is maximized by dissecting thymus lobes in patterns previously shown to be functional for mature T-cell production. The technique is potentially suited to interrogate a milieu of medically relevant questions related to thymus function that include autoimmunity, immunodeficiency and central tolerance; processes which remain mechanistically poorly defined. The fine dissection of mechanisms guiding thymocyte migration, differentiation and selection should lead to novel therapeutic strategies targeting developing T cells.

Real Time In Vivo Tracking of Thymocytes in the Anterior Chamber of the Eye by Laser Scanning Microscopy

The goal of the protocol is to show longitudinal intravital real-time tracking of thymocytes by laser scanning microscopy in thymic implants in the anterior chamber of the mouse eye. The transparency of the cornea and vascularization of the graft allows for continuously recording progenitor cell recruitment and mature T-cell egress.

The purpose of the method being presented is to show, for the first time, the transplant of newborn thymi into the anterior eye chamber of isogenic adult ...