Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Summary

This protocol presents a method for decellularization and subsequent hydrogel formation of murine mammary fat pads following ex vivo irradiation.

Abstract

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue and its role in local recurrence at the primary tumor site are unknown. Here we present a method for the decellularization, lyophilization, and fabrication of ECM hydrogels derived from murine mammary fat pads. Results are presented on the effectiveness of the decellularization process, and rheological parameters were assessed. GFP- and luciferase-labeled breast cancer cells encapsulated in the hydrogels demonstrated an increase in proliferation in irradiated hydrogels. Finally, phalloidin conjugate staining was employed to visualize cytoskeleton organization of encapsulated tumor cells. Our goal is to present a method for fabricating hydrogels for in vitro study that mimic the in vivo breast tissue environment and its response to radiation in order to study tumor cell behavior.

Introduction

Cancer is characterized by excess proliferation of cells that can evade apoptosis and also metastasize to distant sites¹. Breast cancer is one of the most common forms among females in the US, with an estimated 266,000 new cases and 40,000 deaths in 2018². A particularly aggressive and difficult to treat subtype is triple negative breast cancer (TNBC), which lacks estrogen receptor (ER), progesterone receptor (PR), and human epidermal growth factor (HER2). Radiation therapy is commonly used in breast cancer to eliminate residual tumor cells following lumpectomy, but over 13% of TNBC patients still experience recurrence at the primary tumor site³.

It is known that radiation therapy is effective in mitigating metastasis and recurrence because the combination of lumpectomy and radiation results in the same long-term survival as mastectomy⁴. However, it has recently been shown that radiation treatment is associated with local recurrence to the primary tumor site in immunocompromised settings^5,6. Also, it is well known that radiation changes the extracellular matrix (ECM) of normal tissue by inducing fibrosis⁷. Therefore, it is important to understand the role of radiation-induced ECM changes in dictating tumor cell behavior.

Decellularized tissues have been used as in vitro models to study disease^8,9. These decellularized tissues preserve ECM composition and recapitulate the complex in vivo ECM. This decellularized tissue ECM can be further processed and digested to form reconstituted ECM hydrogels that can be used to study cell growth and function^10,11. For example, injectable hydrogels derived from decellularized human lipoaspirate and from myocardial tissue served as non-invasive methods of tissue engineering, and a hydrogel derived from porcine lung tissue was utilized as an in vitro method of testing mesenchymal stem cell attachment and viability^12,13,14. The effect of normal tissue radiation damage on ECM properties, however, has not been investigated.

Hydrogels derived from ECM have the greatest potential for in vitro study of in vivo phenomena. Several other materials have been studied, including collagen, fibrin, and matrigel, but it is difficult to synthetically recapitulate the composition of the ECM¹³. An advantage of using ECM-derived hydrogels is that the ECM contains the necessary proteins and growth factors for a particular tissue^14,15. Irradiation of normal tissue during lumpectomy causes significant changes to the ECM, and ECM-derived hydrogels can be used to study this effect in vitro. This method could lead to more complex and more accurate in vitro models of disease.

In this study, we subjected murine mammary fat pads (MFPs) to radiation ex vivo. The MFPs were decellularized and made into pre-gel solution. Hydrogels were formed with embedded 4T1 cells, a murine TNBC cell line. The rheological properties of the hydrogel material were examined, and tumor cell dynamics were evaluated within the hydrogels. Hydrogels fabricated from irradiated MFPs enhanced tumor cell proliferation. Future studies will incorporate other cell types to study cell-cell interactions in the context of cancer recurrence following therapy.

Protocol

Animal studies were performed in accordance with institutional guidelines and protocols approved by the Vanderbilt University Institutional Animal Care and Use Committee.

1. Preparation and ex vivo irradiation of MFPs

1. Sacrifice athymic Nu/Nu mice (8–10 weeks) using CO­₂ asphyxiation followed by cervical dislocation.
2. Clean the skin using 70% ethanol.
3. Collect mammary fat pads (MFPs) from sacrificed mice using pre-sterilized scissors and forceps in a 15 mL conical tube containing complete RPMI media (RPMI supplemented with 1% penicillin-streptomycin and 10% fetal bovine serum) (see the Table of Materials).
4. Irradiate samples to 20 Gy using a cesium source.
5. Bring the irradiated MFPs and complete RPMI media into a biosafety cabinet. The media will be dependent on the cell line to be grown in the final hydrogel. Fill 6 cm or 10 cm dishes with enough media to submerge the MFPs. For 6 cm dishes, use 8 mL of media, and for 10 cm dishes, use 20 mL of media.
6. Incubate in a 37 °C/5% CO₂ incubator for two days. The length of time in the incubator may be adjusted.
7. Rinse tissues in phosphate-buffered saline (PBS), blot excess moisture, and place MFPs into 15 mL conical tubes for storage at -80 °C until decellularization. This freezing step aids the decellularization step and the sample should be frozen even if the next steps are otherwise ready.

2. Decellularization (adapted from references^12,16,17)

NOTE: This procedure was adapted from previously published methods focused on adipose decellularization, which included the sodium deoxycholate ionic detergent rather than sodium dodecyl sulfate to remove DNA efficiently^12,16,17.

1. On day 1, remove frozen MFPs from -80 °C and thaw at room temperature.
2. Once thawed, dry MFPs briefly on a delicate task wipe. Weigh the MFPs using an analytical scale.
3. Using a pair of forceps with scissors or a scalpel, divide tissue up into 3 mm x 3 mm x 3 mm samples for study of the intact ECM and the remaining tissue for hydrogel production.
   NOTE: The number of samples is dependent on the number of testing methods, e.g. the collection of two samples is described below: one for paraffin embedding (step 2.5) and one for freezing in cryostat embedding medium, if desired (see the Table of Materials and step 2.6).
4. Weigh the tissues. If embedding in paraffin for sectioning, continue to step 2.5. If freezing in cryostat embedding medium for sectioning, continue to step 2.6.
5. In a chemical hood, submerge the tissue in 10% neutral buffered formalin (NBF) (see the Table of Materials) for 24 h at 4 °C. Wash 3 times in PBS for 5 min each. Submerge the tissue in 30% sucrose for 48 h at 4 °C.
   1. Weigh the tissue now that this piece has been removed. Continue to step 2.6.
6. In a chemical hood, place MFP pieces in a labeled cassette prepped with cryostat embedding medium. Add more cryostat embedding medium to cover the tissue.
   1. Place the cassette into a beaker of 2-methylbutane (see the Table of Materials) that is pre-cooled with liquid nitrogen. The beaker should have enough 2-methylbutane to cover the bottom but not enough to submerge the cassette because the cryostat embedding medium should not touch the 2-methylbutane. Let the cassette sit in the 2-methylbutane until the cryostat embedding medium freezes and becomes opaque.
   2. Wrap the cassette(s) in foil, label, and leave at -80 °C until used for sectioning.
      NOTE: Tissues placed immediately in cryostat embedding medium were sectioned at 5 μm while tissues incubated in sucrose were sectioned at 30 μm to retain adipocyte morphology.
7. Use forceps to manually massage the remaining tissue.
   NOTE: Tissue pieces may also be placed in 10% NBF for 24–48 h, rinsed in PBS, and left in 70% ethanol until embedding in paraffin. Following embedding, 5 μm sections can be used for hematoxylin and eosin (H & E) staining (see section 7 below).
8. Place the MFPs in 6 cm dishes with 5 mL 0.02% trypsin/0.05% EDTA solution. Incubate at 37 °C for 1 h. Spray and wipe the dishes with 70% ethanol before placing in the incubator.
9. Use 0.7 mm strainers to wash the MFPs with deionized (DI) water by pouring water over the tissue three times. Use forceps to manually massage the tissue in between washes.
10. Briefly dry tissue on a delicate task wipe and weigh. Place tissues in a pre-autoclaved beaker containing an appropriately sized stir bar. Cover tissues with 60 mL of 3% t-octylphenoxypolyethoxyethanol (see the Table of Materials) per 1 g of tissue and stir for 1 h at room temperature. Use a minimum of 20 mL.
11. Dump tissue and contents into a strainer. Rinse the beaker with DI water and pour onto tissues. Repeat two more times. Use forceps to manually massage the tissue in between rinses.
12. Briefly dry tissue on a delicate task wipe and weigh. Place tissues and stir bars back in the same beakers, and cover with 60 mL of 4% deoxycholic acid per 1 g of tissue. Stir for 1 h at room temperature. Use a minimum of 20 mL.
13. Dump tissue and contents into a mesh strainer. Rinse the beaker with DI water and pour onto tissues. Repeat two more times. Use forceps to manually massage the tissue in between rinses.
14. Briefly dry tissue on a delicate task wipe and weigh.
15. Place tissues in the same beaker with fresh DI water supplemented with 1% penicillin-streptomycin. Cover tightly with paraffin film. Leave overnight at 4 °C.
16. Wash strainers and beakers for use the following day.
17. On day 2, drain beaker contents into a strainer. Briefly dry tissue on a delicate task wipe and weigh.
18. Place MFPs in the same beaker with an appropriately sized stir bar. Cover with 60 mL 4% ethanol/0.1% peracetic acid solution per 1 g of tissue. Use a minimum of 20 mL. Stir for 2 h at room temperature.
19. Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue. Place contents back into the beaker. Wash tissue by covering it with 60 mL of 1x PBS per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat once.
20. Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue. Place contents back into beaker. Wash tissue by covering it with 60 mL DI water per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat once.
21. Briefly dry tissue on a delicate task wipe and weigh. Dump tissue and contents into a strainer. Use forceps to manually massage the tissue.
22. Place contents back into beaker. Cover tissues with 60 mL of 100% n-propanol per 1 g of tissue. Use a minimum of 20 mL. Stir for 1 h at room temperature.
23. Briefly dry tissue on a delicate task wipe and weigh. Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue.
24. Place contents back into beaker. Wash tissue by covering it with 60 mL of DI water per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat three times.
25. Dump tissue and contents into a strainer. Repeat steps 2.3–2.6 to collect pieces of tissue for sucrose incubation and freezing in cryostat embedding medium.
26. Briefly dry tissue on a delicate task wipe and weigh. Place in a labeled 15 mL tube. Freeze at -80 °C overnight.

3. Lyophilization

1. Remove the 15 mL tubes from -80 °C and place on dry ice. Keep samples frozen on dry ice until on the lyophilizer.
2. Remove caps. Use a rubber band to attach a delicate task wipe to the top to cover the opening. Put samples on the lyophilizer for at least 2 days.
3. Remove samples from the lyophilizer and place tubes on dry ice. Remove delicate task wipes weigh each sample on an analytical scale. Attach caps and place at -80 °C overnight.

4. Milling

1. Fill a shallow container with liquid nitrogen. Remove samples from the -80 °C freezer. Weigh each lyophilized MFP.
2. Place one sample in the mortar. Use a cryogenic glove to hold the mortar in the liquid nitrogen.
3. Use a pestle attached to a handheld drill to mill the sample. Mill in 1 min intervals to check progress and remove gloved hand from liquid nitrogen. Mill for a minimum of 5 min.
4. Repeat for all samples. Spray and wipe the mortar and pestle with ethanol between each sample.
5. Store powdered samples in 15 mL tubes at -80 °C until ready for use.
   NOTE: Samples may be used immediately or stored overnight.

5. Hydrogel formation

1. If stored at -80 °C overnight, remove and thaw at room temperature. While thawing, calculate the necessary weight of pepsin (see Table of Materials) and volume of hydrochloric acid (HCl) needed for each sample that results in a solution with 1% w/v sample powder and 0.1% w/v pepsin in 0.01 M HCl. Add the pepsin to the HCl to form a pepsin-HCl solution.
2. Add sample powder and pepsin-HCl solution to a 15 mL tube. Add a small stir bar, and stir for 48 h.
3. Place the tubes on ice for 5 min. Calculate the necessary volume of 10x PBS needed for each sample that results in a solution with a 1x PBS concentration. Add the appropriate volume of 10x PBS to each tube.
4. Add 10% v/v 0.1 M NaOH to each solution to reach pH 7.4. Use a pH paper to test individually.
   NOTE: Gel solution may be stored at 4 °C for one week.

6. Encapsulating cells in hydrogels

1. Using GFP- and luciferase-labeled cells
   1. Using the pH 7.4 gel solution, resuspend pelleted GFP- and luciferase-labeled 4T1 cells to a concentration of 500,000 or 1,000,000 cells/mL of gel solution. Add 16 µL of gel-cell solution to each well of a 16-well chamber slide. Incubate for 30 min at 37 °C.
   2. Add 100 µL of complete RPMI media to each well. Continue incubation for 48 h at 37 °C. The GFP- and luciferase-labeled cells can be visualized using fluorescence microscopy at 0 hours, 24 hours, and 48 hours after gelation.
      NOTE: Cell proliferation can be measured for the GFP- and luciferase-labeled 4T1 cells used here by adding (S)-4,5-Dihydro-2-(6-hydroxy-2-benzothiazolyl)-4-thiazolecarboxylic acid potassium salt (see the Table of Materials) to the wells for 10 min and performing bioluminescence imaging using a bioluminescence imaging system (see Table of Materials). Following bioluminescence imaging, the cytoskeleton can be visualized (see section 10).
2. Using unlabeled cells
   1. Using the pH 7.4 gel solution, resuspend pelleted unlabeled 4T1 cells to a concentration of 500,000 or 1,000,000 cells/mL of gel solution. Add 16 µL of gel-cell solution to each well of a 16-well chamber slide and a 96-well plate. Incubate for 30 min at 37 °C.
   2. Add 100 µL of media to each well. Continue incubation for 48 h at 37 °C.
      NOTE: Cell viability can be measured (see section 11). The 16-well chamber slide can be used for imaging, and the 96-well plate can be used for quantification.

7. H & E staining

1. For formalin-fixed, paraffin-embedded tissue, submerge slides containing 5 μm sections twice in 100% xylene (5–10 min) for de-paraffinization. Rehydrate by submerging in 100%, 95%, 85%, and 70% ethanol for 5 min each followed by DI water for 5 min. Continue to step 7.6.
2. For frozen tissue sections, take sections immediately from freezer and submerge them in 10% NBF for 10 min. Wash in 1x PBS three times for 5 min each. Rinse in water for 5 min.
3. Stain nuclei with hematoxylin for 3 min. Rinse in running tap water.
4. Differentiate by dipping 1–2 times in 0.3% acid alcohol (0.3% HCl in 70% ethanol). Rinse in running tap water for 5 min.
5. Add bluing agent for 30 s. Rinse in running tap water for 5 min. Submerge in 95% ethanol for 30 s.
6. Incubate with eosin for 90 s at room temperature. Dehydrate in 3 changes of 100% ethanol for 5 min each.
7. Submerge twice in 100% xylene for 5 min each. Add distyrene-plasticiser-xylene (DPX0 mounting media to the slide and add a coverslip. Let it cure overnight before imaging.

8. 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining

1. Prepare 1-([4-(Xylylazo)xylyl]azo)-2-naphthol solution.
   1. Add 0.5 g of 1-([4-(Xylylazo)xylyl]azo)-2-naphthol powder to a beaker with 100 mL propylene glycol. Heat to 95–100 °C while stirring for at least 30 min. Prevent the temperature from exceeding 100 °C.
   2. Remove beaker from heat and allow to cool slightly. Pour solution through grade 4 qualitative filter paper to remove any residual particulates.
      NOTE: The solution can be stored at room temperature and should be filtered through a 0.45 µm syringe filter immediately before staining.
2. Stain frozen tissue sections.
   1. Remove slides from the -80 °C freezer and air dry for 30 min. Pre-cool 10% NBF at -20 °C for 30 min in a Coplin jar. Fix the slides at room temperature for 10 min. Wash in DI water 3 times for 5 min each.
   2. Remove excess water using a delicate task wipe before immersing in propylene glycol 2 times for 5 min each. Remove slides from propylene glycol. Do not rinse.
   3. Place slides into syringe filtered Oil Red O staining solution for 3 h at room temperature.
   4. Differentiate by placing slides in 85% propylene glycol for 5 min. Rinse samples in PBS twice for 5 min each.
   5. Stain samples with hematoxylin for 30 s. Wash in running tap water for 5 min and then place the slides in DI water.
   6. Mount samples using aqueous based mounting medium and allow media to dry overnight before imaging.

9. Rheology

1. Bring pre-gel solutions from step 5.4 to the rheometer on ice.
2. Attach a 25 mm arm and plate to the rheometer. Open the rheology software on the computer connected to the rheometer.
3. Perform rotational mapping and determine the zero gap. Raise the arm when complete.
4. Pipette 500 μL of pre-gel on the plate. Lower the arm until the pre-gel solution completely occupies the gap between the plate and the arm. Be conservative because lowering the arm past this point will result in pre-gel solution spilling out of the side.
5. Perform a frequency sweep on the pre-gel solution from 0.1–100 Hz after it has remained at 37 °C for 30 min.
6. Raise the rheometer arm when complete. Wipe liquid with a delicate task wipe. Rinse with DI water and wipe with a delicate task wipe. Repeat steps 9.4–9.6 with additional samples.

10. Phalloidin conjugate staining of F-actin

1. Bring phalloidin conjugate solution to room temperature. Briefly centrifuge at a low speed prior to opening. Aliquot enough solution for the assay, and store the rest at -20 °C.
2. Dilute 1000x phalloidin conjugate stock solution to a 1x working solution by adding 1 μL stock solution to 1 mL 1x PBS + 1% bovine serum albumin (BSA). This makes enough for 10 wells (100 μL/well).
   NOTE: One may use plain 1x PBS, but 1x PBS + BSA is preferred to prevent phalloidin conjugate from sticking to the tube. Do not store this solution after the assay. 1 μL of blue fluorescent dye (see the Table of Materials) working solution may be added to stain for nuclei. Working solution may be made by mixing 1 μL of 20 mM blue fluorescent dye with 11.3 μL 1x PBS.
3. Aspirate media from wells (step 6.1). Rinse wells with 1x PBS.
4. Add 10% NBF. Incubate wells with 10% NBF for 20 min at room temperature. Remove supernatant. Wash 3 times with PBS for 5 min each time.
5. Add 0.1% t-Octylphenoxypolyethoxyethanol in PBS for 5 min. Aspirate and wash 3 times with PBS for 5 min each time.
6. Add 100 μL of working solution from step 10.2 to each well. Incubate for 1 h at room temperature. Aspirate and wash 3 times with PBS for 5 min each time. Leave in PBS at 4 °C.
7. Aspirate and remove wells from the gasket on the chamber slide. Use needle nose tweezers to remove the gasket from the slide. Mount and coverslip samples using aqueous based mounting medium. Allow to cure (5 min).
8. Observe cells under a fluorescence microscope at excitation/emission of 590/618 nm.

11. Viability assay

1. Rinse the cells with Dulbecco’s PBS (DPBS). Add 100 µL of DPBS containing 1 µM calcein AM and 2 µM ethidium homodimer to each well. Incubate for 30 min at room temperature.
2. Image the 16-well chamber slide by fluorescence microscopy. Calcein acetoxymethyl (calcein AM) can be observed at excitation/emission = 494/517 nm. Ethidium homodimer can be observed at excitation/emission = 528/645 nm.
3. Read the 96-well plate on a plate reader using the same wavelengths in step 11.2.

Results

MFPs were decellularized following irradiation using the procedure shown in Figure 1A. MFPs pre-decellularization (Figure 1B) and post-decellularization (Figure 1C) are shown. Decellularization was confirmed using hematoxylin and eosin (H & E) staining, and 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining was used to evaluate lipid content (Figure 2). Rheological properties of the ECM hydrogels were also assessed at 37 ...

Discussion

This method of hydrogel formation is largely dependent on the amount of starting tissue. Murine MFPs are small, and the decellularization process results in a significant reduction of material (Table 1). The process can be repeated with more MFPs to increase final yield. Milling is another important step that may lead to loss of material. Others have shown success with a cryogenic mill, but this protocol is based on milling via a handheld mortar and electric drill with a pestle attachment

Materials

Name	Company	Catalog Number	Comments
10% Neutral Buffered Formalin, Cube with Spigot	VWR	16004-128	-
2-methylbutane	Alfa Aesar	19387	-
AR 2000ex Rheometer	TA Instruments	10D4335	rheometer
Bovine Serum Albumin	Sigma-Aldrich	A1933-25G	-
calcein acetoxymethyl (calcein AM)	Molecular Probes, Inc.	C1430	-
D-Luciferin Firefly, potassium salt	Biosynth Chemistry & Biology	L-8820	(S)-4,5-Dihydro-2-(6-hydroxy-2-benzothiazolyl)-4-thiazolecarboxylic acid potassium salt
DPX Mountant for Histology	Sigma-Aldrich	06522-500ML	-
Dulbecco's phosphate-buffered saline	Gibco	14040133	-
Eosin-Y with Phloxine	Richard-Allan Scientific	71304	eosin
ethidium homodimer	Molecular Probes, Inc.	E1169	ethidium homodimer-1 (EthD-1)
Fetal Bovine Serum	Sigma-Aldrich	F0926-500ML	-
Fisher Healthcare Tissue-Plus O.C.T. Compound	Fisher Scientific	23-730-571	cryostat embedding medium
Fluoromount-G	SouthernBiotech	0100-01	aqueous based mounting medium
FreeZone 4.5	Labconco	7751020	lyophilizer
Hoechst 33342 Solution (20 mM)	Thermo Scientific	62249	blue fluorescent dye
Hydrochloric acid	Sigma-Aldrich	258148-500ML	-
IVIS Lumina III	PerkinElmer	CLS136334	bioluminescence imaging system
Kimtech Science Kimwipes	Kimberly Clark		delicate task wipes
n-Propanol (Peroxide-Free/Sequencing), Fisher BioReagents	Fisher Scientific	BP1130-500	-
Oil Red O	Sigma-Aldrich	O0625-25G	1-([4-(Xylylazo)xylyl]azo)-2-naphthol
OPS Diagnostics CryoGrinder	OPS Diagnostics, LLC	CG-08-02	-
PBS (10X), pH 7.4	Quality Biological, Inc.	119-069-151	Phosphate-buffered saline
Penicillin-Streptomycin	Gibco	15140-122	-
Pepsin from porcine gastric mucosa	Sigma-Aldrich	P6887-5G	pepsin
Peracetic acid	Sigma-Aldrich	77240-100ML	-
Phalloidin-iFluor 594 Reagent (ab176757)	abcam	ab176757	phalloidin conjugate
Propylene glycol	Sigma-Aldrich	W294004-1KG-K	-
Richard-Allan Scientific Signature Series Bluing Reagent	Richard-Allan Scientific	7301L	bluing agent
Richard-Allan Scientific Signature Series Hematoxylin 7211	Richard-Allan Scientific	7211	-
RPMI Medium 1640	Gibco	11875-093	-
Sodium deoxycholate, 98%	Frontier Scientific	JK559522	deoxycholic acid
Sucrose	Sigma-Aldrich	S5016	-
Triton x-100	Sigma-Aldrich	X100-100ML	t-Octylphenoxypolyethoxyethanol
Trypsin-EDTA (0.25%), phenol red	Gibco	25200-056	-
Whatman qualitative filter paper, Grade 4	Whatman	1004-110	grade 4 qualitative filter paper
Xylenes (Certified ACS), Fisher Chemical	Fisher Scientific	X5-4	-