Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

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-catenin^3,4, 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 microscopy^5,6. 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 software^7,10. 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 livers^8,11. 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 software^3,12,13,14. 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.

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

1. 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.
2. 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.
3. 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.
4. Incubate at 4 °C at least overnight (but up to several months) with gentle rocking.

2. Dissecting Tissues Surrounding Liver

1. 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.
2. Pipette several larvae in PBS into one well of a 9-well round-bottom glass dish.
3. Remove skin surrounding liver.
   1. 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.
   2. 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).
   3. Continue grabbing flaps of skin and pulling down/back until all of the skin and melanophores overlying or near the liver have been removed.
4. Remove yolk, if present.
   1. 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.
   2. 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.
5. Place dissected larvae into fresh cold PBS using a glass pipette and pipette pump.

3. Imaging

1. To mount larvae, pour a few mL of 3% methyl cellulose onto the lid of a clean plastic Petri dish.
2. Use a glass pipette and pipette pump to add larvae to the methyl cellulose, adding as little PBS with the larvae as possible.
3. 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.
4. Take a picture of each fish.
   1. 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.
   2. Zoom in to high magnification and focus on the liver, making sure that the liver's outline is clearly visible.
   3. Snap a picture and save the file.
   4. Repeat for all fish, making sure the magnification is the same for each picture.
   5. Take a picture of a micrometer using the same magnification (see Figure 2H).

4. Image Analysis

1. Measure the area of each fish's liver using image processing software.
   1. 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.
   2. Open randomized files in order, starting with file 1.
   3. Choose the freehand selections tool and outline each liver.
   4. Press Ctrl-M to measure the area of each liver.
   5. 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).
   6. Save the measurements in a text file ("measurements file").
2. Un-blind and analyze data
   1. Open "measurements file" and "randomization file" in a spreadsheet program.
   2. 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".
   3. Sort data by original file name.
   4. Be sure to exclude any liver measurements for which pictures were inadequate (see Figure 2A–G).
   5. If necessary, convert measurement values into desired scale (mm², for example).
      1. Open the scale bar in the image processing software.
      2. 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.
      3. Use the conversion factor to convert measurements in the "unblinded measurements file".
   6. Using the spreadsheet program or pasting data into a scientific graphing and statistics software, determine mean and standard deviation and calculate p value(s).

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...

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 pre...

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