Use of Frozen Tissue in the Comet Assay for the Evaluation of DNA Damage

Summary

This protocol describes several procedures for preparing high quality frozen tissue samples at the time of necropsy for use in the comet assay to assess DNA damage: 1) minced tissue, 2) scraped epithelial cells from the gastrointestinal tract, and 3) cubed tissue samples, requiring homogenization using a tissue mincing device.

Abstract

The comet assay is gaining popularity as a means to assess DNA damage in cultured cells and tissues, particularly following exposure to chemicals or other environmental stressors. Use of the comet assay in regulatory testing for genotoxic potential in rodents has been driven by adoption of an Organisation for Economic Co-operation and Development (OECD) test guideline in 2014. Comet assay slides are typically prepared from fresh tissue at the time of necropsy; however, freezing tissue samples can avoid logistical challenges associated with simultaneous preparation of slides from multiple organs per animal and from many animals per study. Freezing also enables shipping samples from the exposure facility to a different laboratory for analysis, and storage of frozen tissue facilitates deferring a decision to generate DNA damage data for a given organ. The alkaline comet assay is useful for detecting exposure-related DNA double- and single-strand breaks, alkali-labile lesions, and strand breaks associated with incomplete DNA excision repair. However, DNA damage can also result from mechanical shearing or improper sample processing procedures, confounding the results of the assay. Reproducibility in collection and processing of tissue samples during necropsies may be difficult to control due to fluctuating laboratory personnel with varying levels of experience in harvesting tissues for the comet assay. Enhancing consistency through refresher training or deployment of mobile units staffed with experienced laboratory personnel is costly and may not always be feasible. To optimize consistent generation of high quality samples for comet assay analysis, a method for homogenizing flash frozen cubes of tissue using a customized tissue mincing device was evaluated. Samples prepared for the comet assay by this method compared favorably in quality to fresh and frozen tissue samples prepared by mincing during necropsy. Moreover, low baseline DNA damage was measured in cells from frozen cubes of tissue following prolonged storage.

Introduction

The comet assay is increasingly used as a means to evaluate DNA damage in cultured cells and tissues exposed to chemicals or other environmental stressors¹. The assay can detect DNA double- and single-strand breaks, alkali-labile lesions, and single-strand breaks associated with incomplete DNA repair. The International Council for Harmonisation of Technical Requirements for Pharmaceuticals for Human Use (ICH) guideline for pharmaceutical testing recommends a DNA strand breakage assay such as the comet assay as a second test to supplement the rodent erythrocyte micronucleus assay for assessing in vivo genotoxicity and as a follow-up test for assessing mode of action in target organs of tumor induction². The European Food Safety Authority (EFSA) recommends the in vivo comet assay as a suitable follow-up test for investigating the relevance of a positive result in an in vitro genotoxicity test³. In 2014, an OECD test guideline was approved for the rodent comet assay, thus increasing the acceptability of the assay for use in regulatory testing of genotoxic potential. The assay is based on the electrophoretic separation of relaxed DNA loops and fragments that migrate from nucleoids of lysed cells. Basically, single cells are embedded in agarose that has been layered onto microscope slides. Slides are then immersed in lysis buffer followed by an alkaline (pH > 13) solution, which allows the tightly coiled nuclear DNA to relax and unwind. Slides are then placed in an electric field, which stimulates migration of negatively charged DNA toward the anode, creating images that resemble comets; the relative amount of DNA in the comet tail compared with the comet head is directly proportional to the amount of DNA damage; DNA content in the tail is typically quantified using digital imaging software.

Because the comet assay detects fragmented DNA, accurate quantification of exposure-induced DNA damage can be confounded by chromatin fragmentation associated with necrosis or apoptosis resulting from treatment-induced cytotoxicity or stress. Furthermore, DNA damage can occur as a result of mechanical shearing or improper sample processing⁴. The importance of maintaining harvested tissue chilled prior to slide preparation to minimize the baseline level of DNA damage has been previously demonstrated^5,6. Many laboratories prepare comet assay slides from fresh tissue; however, this can be logistically challenging when preparing slides from multiple tissue types per animal in a study with a large number of animals. Moreover, this presents a problem when slide preparation and analysis are to occur at a remote laboratory, necessitating shipment of samples. For example, the U.S. National Toxicology Program includes the comet assay as a component of its genetic toxicology testing program (https://ntp.niehs.nih.gov/testing/types/genetic/index.html) and sometimes incorporates the assay into 28 or 90 day repeat dose toxicity studies; this necessitates collection of tissue by the in-life laboratory and transfer of samples to another laboratory for analysis. To accomplish this, tissue pieces are minced, and/or epithelial cells of the gastrointestinal tract are scraped and cellular suspensions are flash frozen and stored in a freezer for subsequent shipment and storage by the receiving laboratory until analysis⁷. Proper handling of samples is crucial for obtaining high quality data using frozen tissue; however, reproducible manipulation of tissue samples during necropsies performed by ever-changing personnel is difficult to control, especially at in-life laboratories that do not routinely harvest tissues for the comet assay. Refresher training of necropsy staff or use of a mobile unit staffed by experienced laboratory personnel to collect fresh or frozen tissue samples is often too costly, not feasible, or simply undervalued.

To better ensure consistent generation of high quality tissue samples for transfer to a remote site for comet assay analysis, the utility of a published method⁶ of tissue preservation from flash frozen cubes of tissue was explored. In this method, frozen cubes of tissue are loaded into a stainless steel tissue mincing device (Figure 1) that is placed into a microcentrifuge tube containing cold buffer. The cube of tissue is then pushed through a small gauge mesh at the end of the device. Repeatedly forcing the tissue suspension through the mesh sieve in both directions several times results in a relatively uniform single cell suspension. Samples prepared by this method compared favorably in quality to both fresh and frozen tissue samples prepared by mincing. As an added benefit, unlike minced samples, tissue cubes can be stored frozen for prolonged periods of time and still yield high quality results in the comet assay.

Protocol

Tissues were harvested during the conduct of studies performed at AAALAC-accredited facilities at NTP contract laboratories in accordance with Good Laboratory Practice regulations (21 CFR Part 58) and animal use protocols approved by the Institutional Animal Care and Use Committee (IACUC) at each laboratory.

1. Tissue Harvest and Processing

NOTE: It is useful to prepare duplicate sample tubes (e.g., liver) or transfer approximately half of a sample to another storage tube (e.g., duodenum, stomach) to enable reanalysis, if necessary. To minimize potential sample-to-sample variability for intestinal tract tissues, it is recommended that care be taken to sample the same region of tissue relative to the stomach for each animal, and a sample be divided to generate duplicate samples. Plastic forceps are recommended for transferring sticky tissues such as brain. As an option, minced, scraped or homogenized tissue preparations may be filtered to achieve homogeneous single cell suspensions using a 40 µm cell strainer attached to a conical tube.

1. Remove any attached connecting tissue, organs, and debris from organ(s) of interest. Immediately following removal, swish the organ vigorously in a medium-sized weigh boat containing ~7 mL of cold freshly prepared mincing solution (Mg⁺⁺, Ca⁺⁺ and phenol red-free Hank’s Balanced Salt Solution, 10% v/v DMSO, and 20 mM EDTA pH 7.5) to remove residual blood and debris.
2. Transfer each organ to another clean weigh boat containing sufficient (~7 mL) cold mincing solution to maintain the tissue submerged, on ice, until further processing.
3. Remove a section of each organ of interest and place it in an embedding cassette. Fix in 10% neutral buffered formalin, trim, and paraffin embed according to the laboratory’s standard procedures for possible future histopathology evaluation.
4. For the liver, cut a 5 mm longitudinal section of the left lobe and gently swish in ~7 mL of cold mincing solution. Place the strip of tissue into a clean medium-sized weigh boat containing ~7 mL of cold mincing solution and maintain on ice until ready to process further (step 1.8 or 1.11).
5. For the duodenum, cut a 10 mm portion of the duodenum proximal to the stomach. Using a 21–25 G needle, flush to remove debris and bacteria. 
   1. Insert needle into one end of the duodenum and flush with 1 mL of cold mincing solution.
   2. Flush the other direction by flipping the duodenum over and repeating with another 1 mL of mincing solution. Discard the needle.
   3. Slice the duodenum open and rinse it in ~7 mL of mincing solution before putting it into a medium-sized weigh boat containing ~7 mL of clean mincing solution; maintain on ice until ready to process further (step 1.8 or 1.11).
6. For the brain, it may be useful to first divide the brain into two hemispheres; one hemisphere can be saved for possible histopathology (see step 1.2).
   1. Dissect the brain region(s) of interest and gently swish in ~7 mL of cold mincing solution.
   2. Transfer tissue(s) to a clean medium-sized weigh boat containing ~7 mL of cold mincing solution and maintain on ice until ready to process further (step 1.8 or 1.11; note that small regions such as the cerebellum and hippocampus may not require any further trimming).
7. For the stomach
   1. Remove and discard the forestomach. Cut open the glandular stomach and wash free from food using ~7 mL of cold mincing buffer in a medium-sized weigh boat.
   2. Remove a 5 mm strip of glandular stomach proximal to the duodenum for fixation for possible histopathology evaluation (see step 1.2).
   3. Place the remaining stomach into ~7 mL of cold mincing buffer in a medium-sized weigh boat and incubate on ice for 15–30 min.
      NOTE: This incubation step may not be necessary; this step was included in the JaCVAM validation protocol, but is not mentioned in the OECD test guideline^8,9.
   4. Transfer the stomach to a clean piece of paraffin (or Petri dish or weigh boat) and gently scrape the surface epithelium two or more times using a scalpel blade or a polytetrafluoroethene (PTFE) scraper to remove debris. Pick up the gastric mucosa with forceps and rinse with 1 mL of cold mincing buffer using a pipet. Transfer the stomach tissue to a clean surface or dish.
   5. Carefully scrape the stomach epithelium 4–5 times (more, if necessary) in mincing solution (typically 250 µL/mouse or 500-1,000 µL/rat) with the back of a scalpel blade or PTFE scraper to release the cells. Optionally, rinse tissue with an approximately equal volume of mincing solution to recover more cells. Collect the mincing solution containing released cells using a pipet and transfer to a microcentrifuge tube on ice.
8. For mincing tissue samples, cut a 3–4 mm section of liver or brain tissue or ~5 mm of duodenum and transfer to a labeled 1.5 or 2.0 mL microcentrifuge tube containing 1 mL of cold mincing solution. Rapidly mince (≤30 s) with mincing scissors until finely dispersed, while keeping the sample cold. The sample should look slightly cloudy; however, it is common for some pieces to remain that will settle to the bottom of the tube.
9. To use the tissue fresh, maintain tubes containing minced tissue or scraped epithelial cells on ice; use the tissue to prepare comet slides within approximately 1 h of harvest (outlined in section 2).
10. For freezing the tissue samples, immediately following mincing or collection of scraped epithelial cells
    1. Secure the tube lid and drop tube into a Dewar flask containing liquid nitrogen; tubes may be maintained in liquid nitrogen for the duration of necropsy (≤3 h).
    2. Retrieve tubes using a ladle and place on dry ice to sort. Transfer tubes to a freezer box on dry ice and store box in a -80 °C freezer.
11. For preparing frozen cubes of tissue, cut several small pieces of tissue (≤4 mm in diameter and ~6 mm in length; Figure 1).
    1. Individually drop them directly into a medium-sized weigh boat or other suitable vessel containing liquid nitrogen.
    2. Wait a few seconds for the pieces to freeze completely and transfer individual frozen tissue cubes to a labeled microcentrifuge tube or cryotube maintained on dry ice; do not allow the pieces to clump together during the freezing process.
    3. Transfer tubes to a freezer box on dry ice and store box in a -80 °C freezer.

2. Slide Preparation

1. For minced/scraped tissue
   1. Maintain tubes containing fresh tissues on wet ice.
   2. Place tubes of frozen tissue into a room temperature water bath until the samples are completely thawed, while ensuring they are kept cold. Immediately transfer the tubes onto ice.
   3. Immediately prior to withdrawing cells for transfer to agarose (step 2.3), mix each cell suspension gently; for non-filtered samples, allow large chunks to settle to the bottom of the tube. Maintain all tubes on ice until slides have been prepared for all samples.
2. For cubed tissue
   1. Retrieve tubes containing tissue cubes from the 80 °C freezer and maintain on dry ice until slide preparation.
   2. Aliquot 1 mL of Merchant’s medium (0.14 M NaCl, 1.47 mM KH₂PO₄, 2.7 mM KCl, 8.1 mM Na₂HPO₄, 10 mM NaEDTA, pH 7.4) or mincing solution into a labeled 1.7 mL microcentrifuge tube for each sample and maintain on ice.
   3. Working rapidly to avoid tissue thawing, place a cube of tissue into the open end of a room temperature tissue mincing device (Figure 1). Multiple tissue pieces may be used; if necessary, the frozen tissue may be cut or cracked into smaller pieces using a cold mortar and pestle to reduce the cube size to fit into the device.
   4. Quickly insert a size-matched syringe plunger into the device to push the tissue to the mesh end which should then be placed into the microcentrifuge tube containing cold mincing solution; avoid forcing the liquid to overflow the tube. Immerse the mesh end of the device for approximately 5–10 s in the cold mincing solution to allow the tissue to thaw completely.
   5. Completely depress the plunger until cells are seen extruding through the mesh pores. Then pull the plunger up approximately 2.5 cm and press down again completely; repeat the process several times until the mincing solution becomes turbid.
   6. If necessary, the sample may be concentrated to increase cell density by refrigerated centrifugation for 5 min at 1100 rpm and removal of a portion of the supernatant.
   7. Repeat the process for all samples using a clean tissue mincing device for each sample.
3. Transfer 50 µL of cell suspension to a tube containing 450 µL of 0.5% low melting point agarose at 37 °C.
   NOTE: Volumes may be adjusted, but the amount of agarose should be ≤1%.
4. Prepare and score slides as described previously^10,11.

Results

Study 1
Liver was harvested from two cohorts of male Sprague Dawley rats administered corn oil for 4 days, staggered by one week. Slides were prepared from freshly minced tissue, frozen minced tissue, and frozen cubed tissue processed in Merchant’s medium or mincing solution using the tissue mincing device. Frozen tissues obtained from animals from the first cohort were evaluated after freezer storage for ~3.5 months. Frozen tissues obtained from animals from the second cohort were evaluated after st...

Discussion

As demonstrated previously^7,12,13, properly handled flash frozen minced tissue provides good results in the comet assay. In fact, baseline % tail DNA values for frozen minced rat and mouse liver prepared in our laboratory are typically ≤6%, as recommended by the OECD test guideline⁹ for freshly minced rat liver samples. Good results have been obtained by multiple laboratories using a variety of tiss...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Cryovials	Corning Costar	430488
Dental Wax Sheets	Electron Microscopy Sciences	72670
Dissecting (Mincing) Micro Scissors	Fisher Scientific	08-953-1B
DMSO	Sigma-Aldrich	D8418
Hank's Balanced Salt Solution	Gibco	14175-079
KCl	Teknova	P0315-10
KH2PO4	Sigma-Aldrich	P9791
Low Melting Point Agarose	Lonza	50081
Microfuge Tubes (1.7 mL)	Corning	3207
Na2EDTA	Sigma-Aldrich	E5134
Na2HPO4	Sigma-Aldrich	S7907
NaCl	Sigma-Aldrich	S6191
Neutral Buffered Formalin	Leica	600
Scalpel Blades	Miltex	4-110
Syringe Plunger (1 mL)	Fisher Scientific or Vitality Medical	14-826-88; 8881901014	Becton Dickinson or Monoject tuberculin syringe
Tissue Mincing Device	NorGenoTech (Oslo, Norway)	None	Small variability in diameter observed which can affect snuggness of plunger.
Tweezers, plastic	Trade Winds Direct	DF8088N	Reinforced nylon, nonsterile, blunt tip, autoclavable; tradewindsdirect.com
Weigh Boats	Krackler Scientific/Heathrow Scientific	6290-14251B