The authors thank Dr. Laura L. Bronsart for providing the GFP- and luciferase-4T1 cells, Dr. Edward L. LaGory for advice on 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining, Dr. Craig L. Duvall for IVIS and lyophilizer use, and Dr. Scott A. Guelcher for rheometer use. This research was financially supported by NIH grant #R00CA201304.

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
<ol>
	<li>Sacrifice athymic Nu/Nu mice (8&#8211;10 weeks) using CO&#173;2 asphyxiation followed by cervical dislocation.</li>
	<li>Clean the skin using 70% ethanol.</li>
	<li>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).</li>
	<li>Irradiate samples to 20 Gy using a cesium source.</li>
	<li>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.</li>
	<li>Incubate in a 37 &#176;C/5% CO2 incubator for two days. The length of time in the incubator may be adjusted.</li>
	<li>Rinse tissues in phosphate-buffered saline (PBS), blot excess moisture, and place MFPs into 15 mL conical tubes for storage at -80 &#176;C until decellularization. This freezing step aids the decellularization step and the sample should be frozen even if the next steps are otherwise ready.</li>
</ol>
2. Decellularization (adapted from references12,16,17)
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 efficiently12,16,17.
<ol>
	<li>On day 1, remove frozen MFPs from -80 &#176;C and thaw at room temperature.</li>
	<li>Once thawed, dry MFPs briefly on a delicate task wipe. Weigh the MFPs using an analytical scale.</li>
	<li>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).</li>
	<li>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.</li>
	<li>In a chemical hood, submerge the tissue in 10% neutral buffered formalin (NBF) (see the Table of Materials) for 24 h at 4 &#176;C. Wash 3 times in PBS for 5 min each. Submerge the tissue in 30% sucrose for 48 h at 4 &#176;C.
	<ol>
		<li>Weigh the tissue now that this piece has been removed. Continue to step 2.6.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Wrap the cassette(s) in foil, label, and leave at -80 &#176;C until used for sectioning. 
		NOTE: Tissues placed immediately in cryostat embedding medium were sectioned at 5 &#956;m while tissues incubated in sucrose were sectioned at 30 &#956;m to retain adipocyte morphology.</li>
	</ol>
	</li>
	<li>Use forceps to manually massage the remaining tissue. 
	NOTE: Tissue pieces may also be placed in 10% NBF for 24&#8211;48 h, rinsed in PBS, and left in 70% ethanol until embedding in paraffin. Following embedding, 5 &#956;m sections can be used for hematoxylin and eosin (H &#38; E) staining (see section 7 below).</li>
	<li>Place the MFPs in 6 cm dishes with 5 mL 0.02% trypsin/0.05% EDTA solution. Incubate at 37 &#176;C for 1 h. Spray and wipe the dishes with 70% ethanol before placing in the incubator.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Dump tissue and contents into a strainer. Rinse the beaker with&#160;DI&#160;water and pour onto tissues. Repeat two more times. Use forceps to manually massage the tissue in between rinses.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh.</li>
	<li>Place tissues in the same beaker with fresh DI water supplemented with 1% penicillin-streptomycin. Cover tightly with paraffin film. Leave overnight at 4 &#176;C.</li>
	<li>Wash strainers and beakers for use the following day.</li>
	<li>On day 2, drain beaker contents into a strainer. Briefly dry tissue on a delicate task wipe and weigh.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Dump tissue and contents into a strainer. Use forceps to manually massage the tissue.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Dump tissue and contents into a strainer. Repeat steps 2.3&#8211;2.6 to collect pieces of tissue for sucrose incubation and freezing in cryostat embedding medium.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Place in a labeled 15 mL tube. Freeze at -80 &#176;C overnight.</li>
</ol>
3. Lyophilization
<ol>
	<li>Remove the 15 mL tubes from -80 &#176;C and place on dry ice. Keep samples frozen on dry ice until on the lyophilizer.</li>
	<li>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.</li>
	<li>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 &#176;C overnight.</li>
</ol>
4. Milling
<ol>
	<li>Fill a shallow container with liquid nitrogen. Remove samples from the -80 &#176;C freezer. Weigh each lyophilized MFP.</li>
	<li>Place one sample in the mortar. Use a cryogenic glove to hold the mortar in the liquid nitrogen.</li>
	<li>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.</li>
	<li>Repeat for all samples. Spray and wipe the mortar and pestle with ethanol between each sample.</li>
	<li>Store powdered samples in 15 mL tubes at -80 &#176;C until ready for use. 
	NOTE: Samples may be used immediately or stored overnight.</li>
</ol>
5. Hydrogel formation
<ol>
	<li>If stored at -80 &#176;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.</li>
	<li>Add sample powder and pepsin-HCl solution to a 15 mL tube. Add a small stir bar, and stir for 48 h.</li>
	<li>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.</li>
	<li>Add 10% v/v 0.1 M NaOH to each solution to reach pH 7.4. Use a pH paper&#160;to test individually. 
	NOTE: Gel solution may be stored at 4 &#176;C for one week.</li>
</ol>
6. Encapsulating cells in hydrogels
<ol>
	<li>Using GFP- and luciferase-labeled cells
	<ol>
		<li>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 &#181;L of gel-cell solution to each well of a 16-well chamber slide. Incubate for 30 min at 37 &#176;C.</li>
		<li>Add 100 &#181;L of complete RPMI media to each well. Continue incubation for 48 h at 37 &#176;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).</li>
	</ol>
	</li>
	<li>Using unlabeled cells
	<ol>
		<li>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 &#181;L of gel-cell solution to each well of a 16-well chamber slide and a 96-well plate. Incubate for 30 min at 37 &#176;C.</li>
		<li>Add 100 &#181;L of media to each well. Continue incubation for 48 h at 37 &#176;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.</li>
	</ol>
	</li>
</ol>
7. H &#38; E staining
<ol>
	<li>For formalin-fixed, paraffin-embedded tissue, submerge slides containing 5 &#956;m sections twice in 100% xylene (5&#8211;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.</li>
	<li>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.</li>
	<li>Stain nuclei with hematoxylin for 3 min. Rinse in running tap water.</li>
	<li>Differentiate by dipping 1&#8211;2 times in 0.3% acid alcohol (0.3% HCl in 70% ethanol). Rinse in running tap water for 5 min.</li>
	<li>Add bluing agent for 30 s. Rinse in running tap water for 5 min. Submerge in 95% ethanol for 30 s.</li>
	<li>Incubate with eosin for 90 s at room temperature. Dehydrate in 3 changes of 100% ethanol for 5 min each.</li>
	<li>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.</li>
</ol>
8. 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining
<ol>
	<li>Prepare 1-([4-(Xylylazo)xylyl]azo)-2-naphthol solution.
	<ol>
		<li>Add 0.5 g of 1-([4-(Xylylazo)xylyl]azo)-2-naphthol powder to a beaker with 100 mL propylene glycol. Heat to 95&#8211;100 &#176;C while stirring for at least 30 min. Prevent the temperature from exceeding 100 &#176;C.</li>
		<li>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 &#181;m syringe filter immediately before staining.</li>
	</ol>
	</li>
	<li>Stain frozen tissue sections.
	<ol>
		<li>Remove slides from the -80 &#176;C freezer and air dry for 30 min. Pre-cool 10% NBF at -20 &#176;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.</li>
		<li>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.</li>
		<li>Place slides into syringe filtered Oil Red O staining solution for 3 h at room temperature.</li>
		<li>Differentiate by placing slides in 85% propylene glycol for 5 min. Rinse samples in PBS twice for 5 min each.</li>
		<li>Stain samples with hematoxylin for 30 s. Wash in running tap water for 5 min and then place the slides in DI water.</li>
		<li>Mount samples using aqueous based mounting medium and allow media to dry overnight before imaging.</li>
	</ol>
	</li>
</ol>
9. Rheology
<ol>
	<li>Bring pre-gel solutions from step 5.4 to the rheometer on ice.</li>
	<li>Attach a 25 mm arm and plate to the rheometer. Open the rheology software on the computer connected to the rheometer.</li>
	<li>Perform rotational mapping and determine the zero gap. Raise the arm when complete.</li>
	<li>Pipette 500 &#956;L 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.</li>
	<li>Perform a frequency sweep on the pre-gel solution from 0.1&#8211;100 Hz after it has remained at 37 &#176;C for 30 min.</li>
	<li>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&#8211;9.6 with additional samples.</li>
</ol>
10. Phalloidin conjugate staining of F-actin
<ol>
	<li>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 &#176;C.</li>
	<li>Dilute 1000x phalloidin conjugate stock solution to a 1x working solution by adding 1 &#956;L stock solution to 1 mL 1x PBS + 1% bovine serum albumin (BSA). This makes enough for 10 wells (100 &#956;L/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 &#956;L 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 &#956;L of 20 mM blue fluorescent dye with 11.3 &#956;L 1x PBS.</li>
	<li>Aspirate media from wells (step 6.1). Rinse wells with 1x PBS.</li>
	<li>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.</li>
	<li>Add 0.1% t-Octylphenoxypolyethoxyethanol in PBS for 5 min. Aspirate and wash 3 times with PBS for 5 min each time.</li>
	<li>Add 100 &#956;L 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 &#176;C.</li>
	<li>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).</li>
	<li>Observe cells under a fluorescence microscope at excitation/emission of 590/618 nm.</li>
</ol>
11. Viability assay
<ol>
	<li>Rinse the cells with Dulbecco&#8217;s PBS (DPBS). Add 100 &#181;L of DPBS containing 1 &#181;M calcein AM and 2 &#181;M ethidium homodimer to each well. Incubate for 30 min at room temperature.</li>
	<li>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.</li>
	<li>Read the 96-well plate on a plate reader using the same wavelengths in step 11.2.</li>
</ol>

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T., Frey, B., Fellner, M., Herrmann, M., Fabry, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrin+5+1+facilitates+cancer+cell+invasion+through+enhanced+contractile+forces.">Integrin 5 1 facilitates cancer cell invasion through enhanced contractile forces.</a> Journal of Cell Science. 124 (3), 369-383 (2011).</li><li>Ahmadzadeh, H., 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=Modeling+the+two-way+feedback+between+contractility+and+matrix+realignment+reveals+a+nonlinear+mode+of+cancer+cell+invasion.">Modeling the two-way feedback between contractility and matrix realignment reveals a nonlinear mode of cancer cell invasion.</a> Proceedings of the National Academy of Sciences. 114 (9), E1617-E1626 (2017).</li></ol>

The authors have nothing to disclose.

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<sup class="xref...

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 settings5,6. 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 disease8,9. 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 function10,11. 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 viability12,13,14. 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 tissue14,15. 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.

<table><tbody><tr><td>10% Neutral Buffered Formalin, Cube with Spigot</td><td>VWR</td><td>16004-128</td><td>-</td></tr><tr><td>2-methylbutane</td><td>Alfa Aesar</td><td>19387</td><td>-</td></tr><tr><td>AR 2000ex Rheometer</td><td>TA Instruments</td><td>10D4335</td><td>rheometer</td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma-Aldrich</td><td>A1933-25G</td><td>-</td></tr><tr><td>calcein acetoxymethyl (calcein AM)</td><td>Molecular Probes, Inc.</td><td>C1430</td><td>-</td></tr><tr><td>D-Luciferin Firefly, potassium salt</td><td>Biosynth Chemistry &amp;amp; Biology</td><td>L-8820</td><td>(S)-4,5-Dihydro-2-(6-hydroxy-2-benzothiazolyl)-4-thiazolecarboxylic acid potassium salt</td></tr><tr><td>DPX Mountant for Histology</td><td>Sigma-Aldrich</td><td>06522-500ML</td><td>-</td></tr><tr><td>Dulbecco&#039;s phosphate-buffered saline</td><td>Gibco</td><td>14040133</td><td>-</td></tr><tr><td>Eosin-Y with Phloxine</td><td>Richard-Allan Scientific</td><td>71304</td><td>eosin</td></tr><tr><td>ethidium homodimer</td><td>Molecular Probes, Inc.</td><td>E1169</td><td>ethidium homodimer-1 (EthD-1)</td></tr><tr><td>Fetal Bovine Serum</td><td>Sigma-Aldrich</td><td>F0926-500ML</td><td>-</td></tr><tr><td>Fisher Healthcare Tissue-Plus O.C.T. Compound</td><td>Fisher Scientific</td><td>23-730-571</td><td>cryostat embedding medium</td></tr><tr><td>Fluoromount-G</td><td>SouthernBiotech</td><td>0100-01</td><td>aqueous based mounting medium</td></tr><tr><td>FreeZone 4.5</td><td>Labconco</td><td>7751020</td><td>lyophilizer</td></tr><tr><td>Hoechst 33342 Solution (20 mM)</td><td>Thermo Scientific</td><td>62249</td><td>blue fluorescent dye</td></tr><tr><td>Hydrochloric acid</td><td>Sigma-Aldrich</td><td>258148-500ML</td><td>-</td></tr><tr><td>IVIS Lumina III</td><td>PerkinElmer</td><td>CLS136334</td><td>bioluminescence imaging system</td></tr><tr><td>Kimtech Science Kimwipes</td><td>Kimberly Clark</td><td></td><td>delicate task wipes</td></tr><tr><td>n-Propanol (Peroxide-Free/Sequencing), Fisher BioReagents</td><td>Fisher Scientific</td><td>BP1130-500</td><td>-</td></tr><tr><td>Oil Red O</td><td>Sigma-Aldrich</td><td>O0625-25G</td><td>1-([4-(Xylylazo)xylyl]azo)-2-naphthol</td></tr><tr><td>OPS Diagnostics CryoGrinder</td><td>OPS Diagnostics, LLC</td><td>CG-08-02</td><td>-</td></tr><tr><td>PBS (10X), pH 7.4</td><td>Quality Biological, Inc.</td><td>119-069-151</td><td>Phosphate-buffered saline</td></tr><tr><td>Penicillin-Streptomycin</td><td>Gibco</td><td>15140-122</td><td>-</td></tr><tr><td>Pepsin from porcine gastric mucosa</td><td>Sigma-Aldrich</td><td>P6887-5G</td><td>pepsin</td></tr><tr><td>Peracetic acid</td><td>Sigma-Aldrich</td><td>77240-100ML</td><td>-</td></tr><tr><td>Phalloidin-iFluor 594 Reagent (ab176757)</td><td>abcam</td><td>ab176757</td><td>phalloidin conjugate</td></tr><tr><td>Propylene glycol</td><td>Sigma-Aldrich</td><td>W294004-1KG-K</td><td>-</td></tr><tr><td>Richard-Allan Scientific Signature Series Bluing Reagent</td><td>Richard-Allan Scientific</td><td>7301L</td><td>bluing agent</td></tr><tr><td>Richard-Allan Scientific Signature Series Hematoxylin 7211</td><td>Richard-Allan Scientific</td><td>7211</td><td>-</td></tr><tr><td>RPMI Medium 1640</td><td>Gibco</td><td>11875-093</td><td>-</td></tr><tr><td>Sodium deoxycholate, 98%</td><td>Frontier Scientific</td><td>JK559522</td><td>deoxycholic acid</td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S5016</td><td>-</td></tr><tr><td>Triton x-100</td><td>Sigma-Aldrich</td><td>X100-100ML</td><td>&lt;em&gt;t&lt;/em&gt;-Octylphenoxypolyethoxyethanol</td></tr><tr><td>Trypsin-EDTA (0.25%), phenol red</td><td>Gibco</td><td>25200-056</td><td>-</td></tr><tr><td>Whatman qualitative filter paper, Grade 4</td><td>Whatman</td><td>1004-110</td><td>grade 4 qualitative filter paper</td></tr><tr><td>Xylenes (Certified ACS), Fisher Chemical</td><td>Fisher Scientific</td><td>X5-4</td><td>-</td></tr></tbody></table>

studying normal tissue radiation effects using extracellular matrix hydrogels

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.

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 &#38; 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 ...

Watch this Scientific Journal Video about Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels at JoVE.com

Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

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

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue ...

studying-normal-tissue-radiation-effects-using-extracellular-matrix

Research

JoVE Journal

Cancer Research

5.8K Views.  Vanderbilt University. This protocol presents a method for decellularization and subsequent hydrogel formation of murine mammary fat pads following ex vivo irradiation.

Video: Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Longitudinal Measurement of Extracellular Matrix Rigidity in 3D Tumor Models Using Particle-tracking Microrheology

The mechanical microenvironment has been shown to act as a crucial regulator of tumor growth behavior and signaling, which is itself remodeled and modified as part of a set of complex, two-way mechanosensitive interactions. While the development of biologically-relevant 3D tumor models have facilitated mechanistic studies on the impact of matrix rheology on tumor growth, the inverse problem of mapping changes in the mechanical environment induced by tumors remains challenging. Here, we describe the implementation of particle-tracking microrheology (PTM) in conjunction with 3D models of pancreatic cancer as part of a robust and viable approach for longitudinally monitoring physical changes in the tumor microenvironment, in situ. The methodology described here integrates a system of preparing in vitro 3D models embedded in a model extracellular matrix (ECM) scaffold of Type I collagen with fluorescently labeled probes uniformly distributed for position- and time-dependent microrheology measurements throughout the specimen. In vitro tumors are plated and probed in parallel conditions using multiwell imaging plates. Drawing on established methods, videos of tracer probe movements are transformed via the Generalized Stokes Einstein Relation (GSER) to report the complex frequency-dependent viscoelastic shear modulus, G*(&#969;). Because this approach is imaging-based, mechanical characterization is also mapped onto large transmitted-light spatial fields to simultaneously report qualitative changes in 3D tumor size and phenotype. Representative results showing contrasting mechanical response in sub-regions associated with localized invasion-induced matrix degradation as well as system calibration, validation data are presented. Undesirable outcomes from common experimental errors and troubleshooting of these issues are also presented. The 96-well 3D culture plating format implemented in this protocol is conducive to correlation of microrheology measurements with therapeutic screening assays or molecular imaging to gain new insights into impact of treatments or biochemical stimuli on the mechanical microenvironment.

Particle-tracking microrheology can be used to non-destructively quantify and spatially map changes in extracellular matrix mechanical properties in 3D tumor models.

The mechanical microenvironment has been shown to act as a crucial regulator of tumor growth behavior and signaling, which is itself remodeled and ...

Bioengineering

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Studying the Effects of Matrix Stiffness on Cellular Function using Acrylamide-based Hydrogels

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and cardiovascular disease1-11. Traditional cell biological approaches to studying cellular function involve culturing cells on a rigid substratum (plastic dishes or glass coverslips) which cannot account for the effect of an elastic ECM or the variations in ECM stiffness between tissues. To model in vivo tissue compliance conditions in vitro, we and others use ECM-coated hydrogels. In our laboratory, the hydrogels are based on polyacrylamide which can mimic the range of tissue compliances seen biologically12. "Reactive" cover slips are generated by incubation with NaOH followed by addition of 3-APTMS. Glutaraldehyde is used to cross-link the 3-APTMS and the polyacrylamide gel. A solution of acrylamide (AC), bis-acrylamide (Bis-AC) and ammonium persulfate is used for the polymerization of the hydrogel. N-hydroxysuccinimide (NHS) is incorporated into the AC solution to crosslink ECM protein to the hydrogel. Following polymerization of the hydrogel, the gel surface is coated with an ECM protein of choice such as fibronectin, vitronectin, collagen, etc.
The stiffness of a hydrogel can be determined by rheology or atomic force microscopy (AFM) and adjusted by varying the percentage of AC and/or bis-AC in the solution12. In this manner, substratum stiffness can be matched to the stiffness of biological tissues which can also be quantified using rheology or AFM. Cells can then be seeded on these hydrogels and cultured based upon the experimental conditions required. Imaging of the cells and their recovery for molecular analysis is straightforward. For this article, we define soft substrata as those having elastic moduli (E) &lt;3000 Pascal and stiff substrata/tissues as those with E &gt;20,000 Pascal.

The effect of substrata stiffness on cellular function can be modeled in vitro using polyacrylamide hydrogels of varying compliances.

Tissue stiffness is an important determinant of cellular function, and changes in tissue stiffness are commonly associated with fibrosis, cancer and ...

Biology

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust mechanical support for in vitro cell culture but lack the necessary protein and ligand composition to elicit physiological behavior from cells. This manuscript describes a fabrication method for a kidney cortex ECM-derived hydrogel with proper mechanical robustness and supportive biochemical composition. The hydrogel is fabricated by mechanically homogenizing and solubilizing decellularized human kidney cortex ECM. The matrix preserves native kidney cortex ECM protein ratios while also enabling gelation to physiological mechanical stiffnesses. The hydrogel serves as a substrate upon which kidney cortex-derived cells can be maintained under physiological conditions. Furthermore, the hydrogel composition can be manipulated to model a diseased environment which enables the future study of kidney diseases.

Here we present a protocol to fabricate a kidney cortex extracellular matrix-derived hydrogel to retain the native kidney extracellular matrix (ECM) structural and biochemical composition. The fabrication process and its applications are described. Finally, a perspective on using this hydrogel to support kidney-specific cellular and tissue regeneration and bioengineering is discussed.

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

Hydrogel Arrays Enable Increased Throughput for Screening Effects of Matrix Components and Therapeutics in 3D Tumor Models

Cell-matrix interactions mediate complex physiological processes through biochemical, mechanical, and geometrical cues, influencing pathological changes and therapeutic responses. Accounting for matrix effects earlier in the drug development pipeline is expected to increase the likelihood of clinical success of novel therapeutics. Biomaterial-based strategies recapitulating specific tissue microenvironments in 3D cell culture exist but integrating these with the 2D culture methods primarily used for drug screening has been challenging. Thus, the protocol presented here details the development of methods for 3D culture within miniaturized biomaterial matrices in a multi-well plate format to facilitate integration with existing drug screening pipelines and conventional assays for cell viability. Since the matrix features critical for preserving clinically relevant phenotypes in cultured cells are expected to be highly tissue- and disease-specific, combinatorial screening of matrix parameters will be necessary to identify appropriate conditions for specific applications. The methods described here use a miniaturized culture format to assess cancer cell responses to orthogonal variation of matrix mechanics and ligand presentation. Specifically, this study demonstrates the use of this platform to investigate the effects of matrix parameters on the responses of patient-derived glioblastoma (GBM) cells to chemotherapy.

The present protocol describes an experimental platform to assess the effects of mechanical and biochemical cues on chemotherapeutic responses of patient-derived glioblastoma cells in 3D matrix-mimetic cultures using a custom-made UV illumination device facilitating high-throughput photocrosslinking of hydrogels with tunable mechanical features.

Cell-matrix interactions mediate complex physiological processes through biochemical, mechanical, and geometrical cues, influencing pathological changes ...

Fabrication Followed by 3D Encapsulation of Tumor Spheroids in PEG Hydrogels

Tumor Spheroid Fabrication and Encapsulation in Polyethylene Glycol Hydrogels for Studying Spheroid-Matrix Interactions

Three-dimensional (3D) encapsulation of spheroids is crucial to adequately replicate the tumor microenvironment for optimal cell growth. Here, we designed an in vitro 3D glioblastoma model for spheroid encapsulation to mimic the tumor extracellular microenvironment. First, we formed square pyramidal microwell molds using polydimethylsiloxane. These microwell molds were then used to fabricate tumor spheroids with tightly controlled sizes from 50-500 &#956;m. Once spheroids were formed, they were harvested and encapsulated in polyethylene glycol (PEG)-based hydrogels. PEG hydrogels are a versatile platform for spheroid encapsulation, as hydrogel properties such as stiffness, degradability, and cell adhesiveness can be tuned independently. Here, we used a representative soft (~8 kPa) hydrogel to encapsulate glioblastoma spheroids. Finally, a method to stain and image spheroids was developed to obtain high-quality images via confocal microscopy. Due to the dense spheroid core and relatively sparse periphery, imaging can be difficult, but using a clearing solution and confocal optical sectioning helps alleviate these imaging difficulties. In summary, we show a method to fabricate uniform spheroids, encapsulate them in PEG hydrogels and perform confocal microscopy on the encapsulated spheroids to study spheroid growth and various cell-matrix interactions.

Author Spotlight: In Vitro Hydrogel Model for Glioblastoma Microenvironment Study

Here, we present a protocol that enables fast, robust, and cheap fabrication of tumor spheroids followed by hydrogel encapsulation. It is widely applicable as it does not require specialized equipment. It would be particularly useful for exploring spheroid-matrix interactions and building in vitro tissue physiology or pathology models.

Three-dimensional (3D) encapsulation of spheroids is crucial to adequately replicate the tumor microenvironment for optimal cell growth.  Here, we ...

Comparative Study of Decellularization Techniques for Bovine Lung ECM Hydrogels

Development and Characterization of Decellularized Lung Extracellular Matrix Hydrogels

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural environment in vitro. However, maintaining the native biochemical content of the ECM, achieving mechanical stability, and comprehending the impact of the decellularization process on the mechanical properties of the ECM hydrogels are challenging. Here, a pipeline for decellularization of bovine lung tissue using two different protocols, downstream characterization of the effectiveness of decellularization, fabrication of reconstituted decellularized lung ECM hydrogels and assessment of their mechanical and cytocompatibility properties were described. Decellularization of the bovine lung was pursued using a physical (freeze-thaw cycles) or chemical (detergent-based) method. Hematoxylin and Eosin staining was performed to validate the decellularization and retention of major ECM components. For the evaluation of residual collagen and sulfated glycosaminoglycan (sGAG) content within the decellularized samples, Sirius red and Alcian blue staining techniques were employed, respectively. Mechanical properties of the decellularized lung ECM hydrogels were characterized by oscillatory rheology. The results suggest that decellularized bovine lung hydrogels can provide a reliable organotypic<span style="font-size: 11pt; font-family: Calibri, sans-serif; color: black; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">&#160;alternative to commercial ECM products by retaining most native ECM components. Furthermore, these findings reveal that the decellularization method of choice significantly affects gelation kinetics as well as the stiffness and viscoelastic properties of resulting hydrogels.

Author Spotlight: Optimizing Bovine Lung Decellularization for Organotypic Hydrogels

The protocol elucidates two distinct decellularization methodologies applied to native bovine pulmonary tissues, providing a comprehensive account of their respective characterizations.

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural ...

Decellularizing Mammary Fat Pads: Deriving Extracellular Matrix Hydrogels from Mammary Fat Pads

Source: Alves et. al., <a href="https://www.jove.com/v/59304/studying-normal-tissue-radiation-effects-using-extracellular-matrix">Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels.</a> J. Vis. Exp. (2019).
This video describes a technique of deriving extracellular matrix hydrogels by decellularizing murine mammary fat pads following ex vivo irradiation&#160;for in vitro studies. This hydrogel helps to mimic the in vivo breast tissue environment post-radiation treatment to study tumor cell behavior.

Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Summary

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Chapters in this video

Longitudinal Measurement of Extracellular Matrix Rigidity in 3D Tumor Models Using Particle-tracking Microrheology

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Studying the Effects of Matrix Stiffness on Cellular Function using Acrylamide-based Hydrogels

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Hydrogel Arrays Enable Increased Throughput for Screening Effects of Matrix Components and Therapeutics in 3D Tumor Models

Tumor Spheroid Fabrication and Encapsulation in Polyethylene Glycol Hydrogels for Studying Spheroid-Matrix Interactions

Development and Characterization of Decellularized Lung Extracellular Matrix Hydrogels

Decellularizing Mammary Fat Pads: Deriving Extracellular Matrix Hydrogels from Mammary Fat Pads