Summary
Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.
Abstract
In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.
Introduction
Hepatocellular carcinoma (HCC) is the most common primary malignancy of the liver1 and the third leading cause of cancer-related mortality2. To better understand mechanisms of hepatocarcinogenesis and identify potential HCC therapeutics, we and others have developed transgenic zebrafish in which hepatocyte-specific expression of oncogenes such as β-catenin3,4, Kras(V12)5,6, Myc7, or Yap18 leads to HCC in adult animals. In these zebrafish, liver enlargement is noted as early as 6 days post fertilization (dpf), providing a facile platform for testing the effects of drugs and genetic alterations on oncogene-driven liver overgrowth. Accurate and precise measurement of larval liver size is essential for determining the effects of these manipulations.
Liver size and shape can be assessed semi-quantitatively in fixed zebrafish larvae by CY3-SA labeling9 or in live zebrafish larvae using hepatocyte-specific fluorescent reporters and fluorescence dissecting microscopy5,6. The latter method is relatively quick, and its lack of precision can be addressed by photographing and measuring the area of each liver using image processing software7,10. However, it can be technically challenging to uniformly position all live larvae in an experiment such that two-dimensional liver area is an accurate representation of liver size. A similar technique for quantifying liver size involves using light sheet fluorescence microscopy to quantify larval liver volume8, which may be more accurate for detecting size differences when the liver is expanded non-uniformly in different dimensions. Fluorescence-activated cell sorting (FACS) can be used to count the number of fluorescently labeled hepatocytes and other liver cell types in larval livers8,11. In this method, larval livers are pooled and dissociated, so information about individual liver size and shape is lost. In combination with another liver size determination method, FACS enables differentiation between increased cell number (hyperplasia) and increased cell size (hypertrophy). All of these methods employ expensive fluorescence technology (microscope or cell sorter) and, except for CY3-SA labeling, require labeling of hepatocytes with a fluorescent reporter.
Here we describe in detail a method for quantifying zebrafish larval liver area using bright-field microscopy and image processing software3,12,13,14. This protocol enables precise quantification of the area of individual livers in situ without the use of fluorescence microscopy. While analyzing liver size, we blind the image identity to reduce investigator bias and improve scientific rigor15.
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Protocol
Animal studies are carried out following procedures approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Utah.
1. Fixing Larvae
- At 3–7 days post fertilization (dpf), euthanize larvae with tricaine methanesulfonate (0.03%) and collect up to 15 larvae in a 2 mL tube using a glass pipette and pipette pump.
- Wash larvae twice with 1 mL of cold (4 °C) 1x phosphate-buffered saline (PBS) on ice. For each wash, remove as much liquid as possible from the tube with a glass pipette and pipette pump, and then add 1 mL of cold PBS to tube.
- Remove as much PBS as possible using a glass pipette and pipette pump, and add 1 mL of cold (4 °C) 4% paraformaldehyde (PFA) in PBS.
CAUTION: PFA is an irritant and suspected carcinogen. Gloves should be worn when handling PFA, and concentrated solutions should be handled in a chemical fume hood. - Incubate at 4 °C at least overnight (but up to several months) with gentle rocking.
2. Dissecting Tissues Surrounding Liver
- Remove larvae from PFA by rinsing 3x with 1 mL of cold (4 °C) PBS and rocking for 5 min in between rinses.
NOTE: It is okay to keep the rinsed larvae in PBS for a day or two at 4 °C. - Pipette several larvae in PBS into one well of a 9-well round-bottom glass dish.
- Remove skin surrounding liver.
- Use fine forceps to hold larva on its back (belly up), gripping on either side of the head as gently as possible. Then use very fine forceps in your other hand to grab the skin just overlying the heart.
- Pull skin down diagonally towards the tail of the fish and the bottom of the dish on the left or right side of the fish. Repeat for other side (right or left side).
- Continue grabbing flaps of skin and pulling down/back until all of the skin and melanophores overlying or near the liver have been removed.
- Remove yolk, if present.
- For 5–6 dpf larvae, lift yolk off in one piece by holding the fish with fine forceps on its back and using the very fine forceps to prod the yolk gently.
- For 3–4 dpf larvae, scrape the yolk off in pieces. Hold the fish with fine forceps on its back and use the very fine forceps to stroke the yolk, starting from the ventral side.
- Place dissected larvae into fresh cold PBS using a glass pipette and pipette pump.
3. Imaging
- To mount larvae, pour a few mL of 3% methyl cellulose onto the lid of a clean plastic Petri dish.
- Use a glass pipette and pipette pump to add larvae to the methyl cellulose, adding as little PBS with the larvae as possible.
- Under a dissecting microscope at low magnification, use fine forceps to orient the fish so they are laying on their right side, facing left.
NOTE: Make sure the fish are oriented perfectly on their side or the liver measurements may not be accurate. - Take a picture of each fish.
- Confirm that the fish to be photographed is aligned perfectly, with one eye directly on top of the other eye. If necessary, use fine forceps to tap lightly on head or tail of fish to adjust orientation. If fish's tail is bent, remove the tail by pinching it forcefully with forceps to remove it so the fish lays flat.
- Zoom in to high magnification and focus on the liver, making sure that the liver's outline is clearly visible.
- Snap a picture and save the file.
- Repeat for all fish, making sure the magnification is the same for each picture.
- Take a picture of a micrometer using the same magnification (see Figure 2H).
4. Image Analysis
- Measure the area of each fish's liver using image processing software.
- Blind all liver pictures to avoid potential investigator bias and promote scientific rigor15. This step can be done manually by another lab member or using a computer program (Supplementary Material). Rename files randomly and create a "randomization file" containing a list of the original file names and corresponding blinded file names.
- Open randomized files in order, starting with file 1.
- Choose the freehand selections tool and outline each liver.
- Press Ctrl-M to measure the area of each liver.
- For any livers that cannot be accurately measured, insert a placeholder measurement (very small or very large, so it can be easily excluded later on).
- Save the measurements in a text file ("measurements file").
- Un-blind and analyze data
- Open "measurements file" and "randomization file" in a spreadsheet program.
- Insert a new column in the "measurements file" and add the original file names for the blinded files, using the "randomization file". Save this file as "unblinded measurements file".
- Sort data by original file name.
- Be sure to exclude any liver measurements for which pictures were inadequate (see Figure 2A–G).
- If necessary, convert measurement values into desired scale (mm2, for example).
- Open the scale bar in the image processing software.
- Use the straight line tool to measure 1 mm on the scale bar. The image processing software will measure in the same units as the livers (pixels), giving a conversion factor.
- Use the conversion factor to convert measurements in the "unblinded measurements file".
- Using the spreadsheet program or pasting data into a scientific graphing and statistics software, determine mean and standard deviation and calculate p value(s).
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Representative Results
Transgenic zebrafish expressing hepatocyte-specific activated β-catenin (Tg(fabp10a:pt-β-cat) zebrafish)3 and non-transgenic control siblings were euthanized at 6 dpf and liver area was quantified using brightfield microscopy and image processing software. Transgenic zebrafish have significantly increased liver size (0.0006 cm2) as compared to their non-transgenic siblings (0.0004 cm2, p < 0.0001; Figure 1).
Figure 1: Liver size analysis of 6 dpf (days post fertilization) zebrafish. (A-B) Representative brightfield image of 6 dpf non-transgenic zebrafish larva, which shows natural position and shape of liver overlying the gut. Liver area has been outlined in (B). Scale bars = 0.1 mm. (C-D) Representative brightfield image of 6 dpf transgenic zebrafish larva expressing hepatocyte-specific activated b-catenin (ABC), showing enlarged liver. Liver area has been outlined in (D). Scale bars = 0.1 mm. (E) Graph showing liver size measurements (mean ± standard deviation) of 6 dpf non-transgenic zebrafish larvae (Non-Tg) and transgenic zebrafish larvae expressing hepatocyte-specific activated b-catenin (ABC). Samples were compared using unpaired t test. ****p < 0.0001. Please click here to view a larger version of this figure.
Figure 2: Examples of inadequate images and micrometer. (A-G) Representative images of larval livers that should be excluded from analysis. Scale bars = 0.1 mm. (A) Larva with skin covering the liver. (B) Larva with yolk obscuring the liver. (C) Larva with parts of the liver pinched off. (D) Larva with liver dislocated and falling off. (E) Larva with missing liver. (F) Larva with liver outline that is difficult to identify because image is blurred/out-of-focus. (G) Larva with improper positioning. The two eyes are not aligned directly on top of each other. (H) Image of the micrometer, used to generate scale bars and convert image processing software measurements from pixels to cm2. Please click here to view a larger version of this figure.
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Discussion
Quantification of liver size is crucial in studies aimed at understanding liver development, regeneration, and oncogenesis. The protocol described here is a relatively quick, easy, and cheap technique for liver size quantification in larval zebrafish. Exercising appropriate caution while performing certain aspects of the protocol can aid in increased accuracy of results and decreased frustration.
Proper fixation of the larvae is crucial towards getting well-preserved biological samples and preventing their disintegration. Dilution of the 4% PFA solution can occur when PBS is not removed completely before the addition of PFA to the rinsed larvae. Using well-made PFA solutions and pipetting out all or most of the PBS solution prior to PFA addition is helpful to address this issue.
Although fast and easy to perform after much practice, the dissection technique requires substantial manual dexterity. While dissecting, it is crucial to remove the skin and yolk completely off from above the liver such that the whole liver is exposed. Failure to do this can result in images where the view of the liver boundary is obscured (Figure 2A,B). Unskilled and forceful movements while dissecting can lead to pinching off of parts of the liver (Figure 2C) or loosening of liver attachments, resulting in the liver being displaced (Figure 2D) or missing entirely (Figure 2E). Users should put in adequate numbers of hours towards honing their dissecting skills on practice samples before moving on to experimental samples.
During mounting, the skin above the liver has been removed, increasing the probability of the liver falling out during subsequent steps. To avoid that possibility, gentle pipetting movements should be employed during this process.
During image procurement using the brightfield microscope, it is crucial that good quality images are taken. Blurry, out-of-focus images will make it difficult to assess the true boundary of the liver (Figure 2F). As this method involves measuring the surface area of the left lobe of the liver, it is crucial that the larva is oriented well on its side and not tilted (Figure 2G). Make sure that both eyes of the larva are aligned (one eye covering the view of the other). While measuring surface area using image processing software, it is important to draw the boundary as close as possible to the real outline of the liver so as to avoid measurement discrepancies. Exclude any images where the liver cannot be accurately measured (Figure 2A–G). However, keep in mind that excluding livers can skew the data, as bigger livers are more likely to be disrupted than smaller livers.
One of the limitations of this protocol is that it applies only to fixed larvae. Alternative methods such as fluorescence microscopy can be used to measure liver size in live larvae expressing hepatocyte-specific fluorescent reporters5,7,10. These alternative methods enable sequential measurements to be made on the same animal, and they are also quicker, since they do not require fixation or dissection of the tissues overlying the liver. The advantages of this protocol compared to fluorescence microscopy in live animals are: 1) more flexibility with respect to when livers are measured, as zebrafish can be kept in fixative for weeks or months before photographing them; 2) no requirement for incorporating a fluorescent reporter, which can be cumbersome when dealing with homozygous mutants; and 3) applicability of steps 1 and 2 for other experiments, including immunofluorescence staining or in situ hybridization studies. We use both methods, depending on the particular application. For example, we typically use live imaging and hepatocyte-specific reporters for high-throughput screening3, and follow up on potential hit compounds using the protocol described here3.
This protocol takes only the surface area into account for quantification of liver size, so it does not detect changes in cell metabolism or morphology, nor does it differentiate between increases in cell number and increases in cell size. In order to address this limitation, complementary assays to assess steatosis16, histology6, cell number8,11, cell size3,17, proliferation3,18, and/or apoptosis19 can be performed.
Another limitation of this protocol is that it assumes that increases or decreases in the surface area of the left liver lobe are reflective of the changes in surface area and volume of liver as a whole. This assumption may not apply when liver growth is non-uniform. To examine liver shape and check for non-symmetric increases in liver growth, we routinely do light sheet fluorescence microscopy8 or confocal microscopy3 on our transgenic models. Light sheet fluorescence microscopy can be used to directly quantify larval liver volume8. In transgenic zebrafish expressing hepatocyte-specific Yap1, liver area and liver volume were similarly increased compared to non-transgenic control siblings8.
The dissection technique described here can be combined with immunofluorescence staining, cell-specific fluorescent reporter lines, and/or other labeling techniques to study other aspects of liver development besides liver size3,19,20. As this dissection protocol also exposes the gut and pancreas, it may be helpful for studies of other visceral organs as well.
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Disclosures
The authors have nothing to disclose.
Acknowledgments
We would like to acknowledge Maurine Hobbs and the Centralized Zebrafish Animal Resource (CZAR) at the University of Utah for providing zebrafish husbandry, laboratory space, and equipment to carry out portions of this research. Expansion of the CZAR is supported in part by NIH grant # 1G20OD018369-01. We would also like to thank Rodney Stewart, Chloe Lim, Lance Graham, Cody James, Garrett Nickum, and the Huntsman Cancer Institute (HCI) Zebrafish Facility for zebrafish care. We would like to thank Kenneth Kompass for help with R programming. This work was funded in part by grants from the Huntsman Cancer Foundation (in conjunction with grant P30 CA042014 awarded to Huntsman Cancer Institute) (KJE) and NIH/NCI R01CA222570 (KJE).
Materials
| Name | Company | Catalog Number | Comments |
| Camera for dissecting microscope | Leica, for example | ||
| Dissecting microscope | Leica, for example | ||
| Fine (Dumont #5) forceps | Fine Science Tools | 11254-20 | |
| Glass pipets | VWR | 14672-608 | |
| Image analysis software | Image J/FIJI | ImageJ/FIJI can be dowloaded for free: https://imagej.net/Welcome | |
| Methyl cellulose | Sigma | M0387 | |
| Paraformaldehyde | Sigma Aldrich | P6148 | |
| Phosphate-buffered saline | Various suppliers | ||
| Pipette pump | VWR | 53502-233 | |
| Plastic Petri dishes | USA Scientific Inc | 2906 | |
| Pyrex 9-well round-bottom glass dish | VWR | 89090-482 | |
| Software for blinding files | R project | R can be downloaded for free: https://www.r-project.org/ | |
| Scientific graphing and statistics software | GraphPad Prism | ||
| Spreadsheet program | Microsoft Excel | ||
| Tricaine methanesulfonate (Tricaine-S) | Western Chemical | 200-226 | |
| Very fine (Dumont #55) forceps | Fine Science Tools | 11255-20 |
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